Detection of binding factors with fluorescence polarization

ABSTRACT

This invention relates to a simple and quick method for the detection, identification and/or quantitation of binding factors using fluorescence techniques. A fluorescent probe is incubated with a factor or group of factors, and the presence of a factor capable of binding the probe can be detected by fluorescence polarization. When coupled with a separation step, this invention allows on-line monitoring of binding complex formation.

RELATED APPLICATIONS

[0001] This application claims the benefit of U.S. application Ser. No.60/230,060, filed Sep. 1, 2000, which is hereby incorporated byreference in its entirety.

FIELD OF THE INVENTION

[0002] This invention relates to methods of detection, identificationand quantitation of binding factors with fluorescence techniques.

REFERENCES

[0003] U.S. Pat. No. 6,132,968.

[0004] Baker, D. R., Capillary Electrophoresis, John Wiley & Sons: NewYork; 1995; Chapter 2.

[0005] Bandyopadhyay; P. K., et al., Biochemistry (1978), 17, 4078-4085.

[0006] Barrett, C. H., in Antibody Techniques; Malik, V. S.; Lillehoj,E. P., Eds.; Academic Press: San Diego, 1994; pp 71-102.

[0007] Booth, E. D., et al., (1994) Carcinogenesis 15, 2099-2106.

[0008] Brinkley, M., Bioconjugate Chem. (1992), 3, 2-13.

[0009] Carey, J. Proc. Natl. Acad. Sci. USA (1988), 85, 975-979.

[0010] Chase, J. W., et al., Annu. Rev. Biochem. (1986), 55, 103-136.

[0011] Chen, F. T. et al., Electrophoresis 15 (1994) 13-21.

[0012] Chiem, N. H., et al., Clin. Chem. 44 (1998) 591-598.

[0013] Chu, Y.-H., et al., Acc. Chem. Res. (1995), 28, 461-468.

[0014] Cosman, M., et al., (1990) Carcinogenesis 11, 1667-1672.

[0015] Craig, D. B., et al., Anal. Chem. (1998), 70, 2493-2494.

[0016] Crawford, I. P., et al., Ann. Rev. Biochem. (1980), 49, 163-95.

[0017] Dandliker, W. B., et al., Immunochemistry (1970) 7, 799-828.

[0018] Evangelista, R. A., et al., J. Chromatogr. A 680 (1994) 587-591.

[0019] Fey, H., et al., J. Clin. Microbiol. (1984), 19, 34-38.

[0020] Funk, M., et al., (1997) Bioconjugate Chem 8, 310-317.

[0021] German, I., et al., Anal. Chem. (1998), 70, 4540-4545.

[0022] Gottfried, D. S., et al., J. Phys. Chem. B (1999), 103,2803-2807.

[0023] Guo, X.-Q., et al., Anal. Chem. (1998), 70, 632-637.

[0024] Hamdan, I. I., et al., Nucleic Acids Res. (1998), 26, 3053-3058.

[0025] Haugland, R. P. Handbook of Fluorescent Probes and ResearchChemicals; 6th Edition; Molecular Probes: Eugene, 1996; p 20.

[0026] Hsu, T. M., et al., (1995) Carcinogenesis 16, 2263-2265.

[0027] Johnson, H. M., et al., Appl. Microbiol. (1973), 26, 309-313.

[0028] Karger, B. L. et al., J. Chromatogr. 492 (1989) 585-614.

[0029] Krauss, G., et al., Biochemstry (1981), 20, 5346-5352.

[0030] Lakowicz, J. R., Principles of Fluorescence Spectroscopy; PlenumPress: New York, 1983.

[0031] Lakowicz, J. R. Principles of Fluorescence Spectroscopy; KluwerAcademic/Plenum: New York, 2nd Ed., 1999.

[0032] Lam, M. T., et al., (1999) J Chromatogr A 853, 545-553.

[0033] Lawson, C. L., et al., Nature (1988), 366, 178-182.

[0034] Le, X. C., et al., (1998) Science 280, 1066-1069.

[0035] LeTilly, V., et al., Biochemistry (1993), 32, 7753-7758.

[0036] Lee, M. H., et al., J. Clin. Microbiol. (1987), 25, 1717-1721.

[0037] Lohman, T. M., et al., Annu. Rev. Biochem. (1994), 63, 527-570.

[0038] Margulis, L. A., et al., (1993) Chem Res Toxicol 6, 59-63.

[0039] Marrack, P., et al., Science (1990), 248, 705-711.

[0040] Molineux, I. J., et al., Nucleic Acids Res. (1975), 2, 1821-1837.

[0041] Motulsky, H., Analyzing Data with GraphPad Prism, GraphPadSoftware: San Diego, Calif., 1999; p 173.

[0042] Nix, B.; Wild, D., in “Immunoassay” (edited by Gosling, P. J.),Oxford University Press, 2000. p 246.

[0043] Otwinowski, Z., et al., Nature (1988), 335, 321-329.

[0044] Perrin, F. J., Phys. Radium (1926), 7, 390-401.

[0045] Pfeifer, G. P., (Editor), Technologies for Detection of DNADamage and Mutations, Plenum Press, New York, 1996.

[0046] Santella, R. M., et al., Carcinogenesis 15 (1984) 373-377.

[0047] Scatchard, G., Ann. NY Acad. Sci. USA, (1949, 51, 660.

[0048] Schantz, E. J., et al., Biochemistry (1972), 11, 360-366.

[0049] Schmalzing, D., et al., Anal. Chem. 67 (1995) 606-612.

[0050] Schultz, N. M., et al., Anal. Chem. (1993), 65, 3161-3165.

[0051] Schulz, N. M., et al., Anal. Chem. 67 (1995) 924-929.

[0052] Schwenzer, K. S., et al., Ther. Drug Monit. (1983), 5, 341-345.

[0053] Shimura, K., et al., Anal. Chem. (1994), 66, 9-15.

[0054] Stebbins, M. A , et al., J. Chromatogr. B (1996), 683, 77-84.

[0055] Stebbins, M. A., et al., J. Chromatogr., B (1996), 683,3053-3058.

[0056] Tan, W. G., et al., (2001) J. Chromatogr. A 924, 377-386.

[0057] Tao, L., et al., (1996) Anal. Chem. 68, 3899-3906.

[0058] Thompson, N. E., et al., Appl. Environ. Microbiol. (1986), 51,885-890.

[0059] Wan, Q.-H., et al., Anal. Chem. (1999), 71, 4183.

[0060] Wan, Q. H., et al., (1999) J. Chromatogr. A 853, 555-562.

[0061] Wang, H., et al., (2001) Submitted to Anal. Chem.

[0062] Weber, G. Adv. Protein Chem. (1953), 8, 415-459.

[0063] Xing, J. Z., et al., (2001) Methods in Molecular Biology 162,419-428.

[0064] Ye, L., et al., J. Chromatogr. B (1998), 714, 59-67.

[0065] Ye, L., et al., (1998) J. Chromatogr. B 714, 59-67.

[0066] Zhang, H , et al., Mol. Biol. (1994), 238, 592-614.

[0067] All of the above publications, patents and patent applicationsare herein incorporated by reference in their entirety to the sameextent as if the disclosure of each individual publication, patentapplication or patent was specifically and individually indicated to beincorporated by reference in its entirety.

BACKGROUND OF THE INVENTION

[0068] Affinity binding complex formation is an essential step inbiological or pharmaceutical phenomena. For example, the binding ofproteins to DNA underlines many cellular activities including thecontrol of gene expression, site-specific recombination, replication andrepair of DNA damage. Enzyme-substrate interactions involve therecognition and binding of substrate by the enzyme as the first step.Hormones, neurotransmitters, lymphokines and other effector moleculesbind to their receptors to initiate the cellular process whichultimately lead to achievement of their functions.

[0069] Consequently, affinity binding complexes are also important toolsin biological or pharmaceutical research. For example, drug discoveryoften involves identification of binding factors of a particular targetwhich mediates a disease. A variety of methods have been employed todetect affinity binding complex formation in order to identify thebinding factors. For example, the gel electrophoresis mobility shiftassay (EMSA) is the most commonly used method in the study ofprotein-DNA interactions. This method is based on the observation thatbinding of a protein to DNA fragments leads to a reduction in theelectrophoretic mobility of the DNA fragment in non-denaturingpolyacrylamide or agarose gels. While used extensively, EMSA requiresrelatively large amounts of sample and lengthy analysis time. Moreover,the assay is not suitable when dissociation of protein-DNA complexoccurs during gel electrophoresis.

[0070] As another example, capillary electrophoresis (CE) combined withaffinity recognition has gained a tremendous growth in recent years,with increasing biochemical, clinical, and pharmaceutical applications.A key element of the technique is the use of a molecular recognitionagent, typically a protein that binds to a target molecule with highspecificity and affinity. The complex formation can occur either beforeor during the electrophoretic separation, depending on the stability ofthe resultant complex. In applications such as CE-based immunoassays,however, tight binding of the analyte to the protein is essential toachieve a high degree of sensitivity and reproducibility. Ideally, theaffinity complex thus formed should remain intact throughout theelectrophoretic separation.

[0071] In current practice of affinity CE, the formation and stabilityof the complex are usually established by titration experiments, inwhich a series of solutions containing the substrate and its bindingprotein in various ratios are analyzed. The emergence of a new peak uponaddition of the binding protein to the substrate is taken as theevidence for complex formation and the relative intensitiescorresponding to the complex and the free substrate are used forquantitation. The titration experiments have proved to be very useful instudies of binding interactions; however, they are time-consuming andunable to provide unequivocal identification of the complex when thecomplex is not well separated from the unbound molecules. Therefore,there remains a need for a simple and sensitive method to detectaffinity complex formation.

SUMMARY OF THE INVENTION

[0072] This invention is directed to a simple method based onlaser-induced fluorescence polarization (LIFP) detection of an affinitycomplex of a fluorescent probe and its binding factor. The affinitycomplexes are readily distinguished from the unbound molecules on thebasis of their fluorescence polarization, which is sensitive to changesin the rotational diffusion characteristics arising from molecularassociation or dissociation. The relative increase in fluorescencepolarization upon complex formation varied with the molecular size ofthe binding pairs. A small molecule rotates fast in solution andexhibits a low value of polarization whereas a large molecule exhibits ahigher polarization because of its slower motion under the sameconditions. Thus, changes in fluorescence polarization can reflect theassociation or dissociation status between molecules of interest. Whencombined with capillary electrophoresis or other suitable separationprocedures, this method allows for on-line monitoring of affinitycomplex formation.

[0073] Accordingly, an aspect of this invention is directed to a methodfor detecting a binding factor for a probe, comprising:

[0074] (a) labeling the probe with a fluorophore;

[0075] (b) incubating the labeled probe with a factor or a group offactors which may bind the labeled probe to form a binding complex;

[0076] (c) separating the binding complex and the free probe intodifferent fractions; and

[0077] (d) subjecting each fraction from step (c) to fluorescencepolarization measurement under conditions wherein the binding complexproduces a fluorescence pattern different from that of the free probe,thereby allowing detection of the binding complex.

[0078] The separation step may be performed simultaneously with, orprior to, the fluorescence polarization detection step. The free probeand the bound probe may be separated by any method which is compatiblewith fluorescence polarization. Preferably, the separation method iscapable of being performed in liquid phase. The separation method ismore preferably liquid chromatography or electrokinetic chromatography,and most preferable capillary electrophoresis or capillary gelelectrophoresis.

[0079] In another aspect of the present invention, this method can beapplied to screen a chemical compound library, such as combinatoriallibrary. Thus, in order to identify a compound which is capable ofbinding to a molecule of interest, the molecule of interest is used as aprobe and labeled with a fluorophore. The labeled probe is thenincubated with a compound library and the whole mixture can be analyzedby fluorescence polarization. Alternatively, the mixture is separatedwith a suitable method. The fractions are monitored on-line withfluorescence polarization which can distinguish the free probe from thebound probe, thereby identify whether there is a binding compound in theparticular fraction.

[0080] Similarly, this method can also be applied to screen a mixture ofnatural products, such as a cell lysate or a homogenate of tissue. Thenatural products may come from any source, including animals, plants andmicroorganisms.

[0081] In another aspect of this invention, the method can be used todetermine if a particular sequence or modification exists in a DNA. Forexample, exposure to certain carcinogens induce alkylation or otherkinds of modification of DNA, which may result in mutations and abnormalgene expression to cause cancer formation. By using a specific probewhich binds to a certain DNA sequence or modification, one can detect ifthis sequence or modification exists in the genomic DNA for diagnosispurposes.

[0082] The probe can be a protein (including peptides), particularlyantibodies, enzymes and cell surface receptors. The probe can also be anucleic acid, carbohydrate or carbohydrate derivatives, small organic orinorganic compounds, or any molecule which can be labeled with afluorophore or is naturally fluorescent. The probe is preferably lessthan about 15,000 daltons in molecular weight, more preferably less thanabout 10,000 daltons, yet more preferably less than about 5,000 daltons,still more preferably less than about 3,000 daltons, and most preferablyless than about 1,500 daltons.

[0083] In yet another aspect, this invention provides a method todetermine the binding affinity or stoichiometry of an affinity complex.

[0084] In addition, this invention can also be used to monitor theformation of fluorescently labeled molecules, which may be used asprobes in the present method but are not limited to such use. Labelingof molecules with a fluorophore is often required for high sensitivityquantitation and detection of the molecules. However, it is difficult toidentify whether the desired molecule is labeled or what the labelingefficiency is. This invention provides a fast method to monitor thelabeling process since the labeled molecule can be readilydifferentiated from the free fluorephore because of their difference influorescence polarization.

BRIEF DESCRIPTION OF THE DRAWING

[0085]FIG. 1

[0086] Electropherograms showing vertically and horizontally polarizedfluorescence of the complex formed between fluorescein-labeledvancomycin and its antibody. A 35 cm long, 20 mm i.d., fused silicacapillary was used for separation with a 25 mM disodium tetraborate (pH9.1) as the running buffer. The separation voltage was 25 kV.Fluorescence detection was post-capillary with vertically polarizedexcitation at 488 nm. Iv and Ih correspond to vertically andhorizontally polarized fluorescence intensity, respectively, measured at515 nm.

DETAILED DESCRIPTION OF THE INVENTION

[0087] This invention relates to a simple and quick method for thedetection, identification and/or quantitation of binding factors usingfluorescence techniques. A fluorescent probe is incubated with a factoror group of factors, and the presence of a factor capable of binding theprobe can be detected by fluorescence polarization. When coupled with aseparation step, this invention allows on-line monitoring of bindingcomplex formation.

[0088] Prior to describing the invention in further detail, the termsused in this application are defined as follows unless otherwiseindicated.

[0089] Definitions

[0090] As used herein, a “probe” can be any molecule or substance forwhich it is desired to find a binding factor. Examples of a probeinclude proteins, peptides, nucleic acids, carbohydrates orcarbohydrates derivative, and small organic or inorganic compounds.

[0091] As used herein, a “fluorophore” is any fluorescent substance, forexample, fluorescein.

[0092] As used herein, “a factor or a group of factors” means asubstance or a mixture of substance of any nature. A factor may be aprotein, a peptide, a nucleic acid, a carbohydrate or carbohydratederivative or any other organic or inorganic compound. A group offactors may be a mixture of factors of the same nature, or a mixture offactors of different natures. For example, a cell lysate or homogenateis a group of factors which contains proteins, carbohydrates, lipids,nucleic acids, and any other substance contained in a cell or cells.

[0093] As used herein, a nucleic acid “modification” refers to anychange in the structure of the nucleic acid sequence. Changes in thestructure of a nucleic acid sequence include changes in the covalent andnon-covalent bonds in the nucleic acid sequence. Illustrative of thesechanges are mutations, mismatches, strand breaks, as well as covalentand non-covalent interactions between a nucleic acid sequence, whichcontains unmodified and/or modified nucleic acids, and other molecules.Illustrative of a covalent interaction between a nucleic acid sequenceand another molecule are changes to a nucleotide base (e.g., formationof thymine glycol) and covalent cross-links between double-stranded DNAsequences which are introduced by ultraviolet radiation or bycis-platinum. Yet another example of a covalent interaction between anucleic acid sequence and another molecule includes covalent binding oftwo nucleic acid sequences to psoralen following ultravioletirradiation. Non-covalent interactions between a nucleic acid sequenceand another molecule include non-covalent interactions of a nucleic acidsequence with a molecule other than a nucleic acid sequence and otherthan a polypeptide sequence. Non-covalent interactions between a nucleicacid sequence with a molecule other than a nucleic acid sequence andother than a polypeptide sequence are illustrated by non-covalentintercalation of ethidium bromide or of psoralen between the two strandsof a double-stranded deoxyribnucleic acid sequence.

[0094] As used herein, the term “mutation” refers to a deletion,insertion, or substitution. A “deletion” is defined as a change in anucleic acid sequence in which one or more nucleotides is absent. An“insertion” or “addition” is that change in a nucleic acid sequencewhich has resulted in the addition of one or more nucleotides. A“substitution” results from the replacement of one or more nucleotidesby a molecule which is different molecule from the replaced one or morenucleotides. For example, a nucleic acid may be replaced by a differentnucleic acid as exemplified by replacement of a thymine by a cytosine,adenine, guanine, or uridine. Alternatively, a nucleic acid may bereplaced by a modified nucleic acid as exemplified by replacement of athymine by thymine glycol.

[0095] The term “mismatch” refers to a non-covalent interaction betweentwo nucleic acids, each nucleic acid residing on a different nucleicacid sequence, which does not follow the base-pairing rules. Forexample, for the partially complementary sequences 5′-AGT-3′ and5′-AAT-3′, a G-A mismatch is present.

[0096] The term “strand break” when made in reference to a doublestranded nucleic acid sequence includes a single-strand break and/or adouble-strand break. A single-strand break refers to an interruption inone of the two strands of the double stranded nucleic acid sequence.This is in contrast to a double-strand break which refers to aninterruption in both strands of the double stranded nucleic acidsequence. Strand breaks may be introduced into a double stranded nucleicacid sequence either directly (e.g., by ionizing radiation) orindirectly (e.g., by enzymatic incision at a nucleic acid base).

[0097] As used herein, a “sample” may be a biological sample or anenvironmental sample. Environmental samples include material from theenvironment such as soil and water. Biological samples may be animal(e.g., human), fluid (e.g., blood, plasma and serum), solid (e.g.,stool), tissue, liquid foods (e.g., milk), and solid foods (e.g.,vegetables). A biological sample may comprise a cell, tissue extract,body fluid, chromosomes or extrachromosomal elements isolated from acell, genomic DNA, RNA, cDNA and the like.

[0098] Methods

[0099] In the present invention, fluorescence polarization is used todistinguish between a fluorescently-labeled probe and a complexcontaining both the probe and a factor which binds the probe. Thecomplex exhibits higher polarization than the probe because a smallmolecule, such as the probe, rotates freely in solution and tends toyield no polarization. However, when the probe is bound by anotherfactor and the size of the complex is significantly larger than the freeprobe, the complex rotates much less freely, resulting in significantlyhigher polarization.

[0100] Accordingly, the molecular weight of the probe useful in thepresent invention should be less than about 20,000 daltons. A largermolecule will generate sizable polarization, making any increase inpolarization more difficult to detect. Furthermore, since the sizeincrease upon forming a complex to such a large probe is relatively lesssignificant, it is also harder to further increase the polarization.Preferably, the probe has a molecular weight of less than 15,000daltons. The molecular weight is more preferably less than about 10,000daltons, yet more preferably less than about 5,000 daltons, still morepreferably less than about 3,000 daltons, and most preferably less thanabout 1,500 daltons.

[0101] The probe can be a protein, nucleic acid, carbohydrate, lipid, orany molecule which can be labeled with a fluorophore. In particular,peptides, oligonucleotides and oligosaccharides are good probes due totheir small sizes and easy synthesis. If an antibody is a candidate fora probe, it is preferable to use a fragment of the antibody, such as anFab fragment, rather than the entire antibody.

[0102] The present method can be practiced with or without separation ofthe binding complex and the probe. Thus, the entire incubation mixturecan be subjected to fluorescence polarization without separation, andthe polarization pattern is compared to that of the probe alone. Anincrease in polarization would indicate the presence of at least onebinding factor in the sample used to bind the probe.

[0103] Alternatively, the incubation mixture can be separated accordingto any method known in the art, and then subjected to polarization. Inthis case, the free probe and the complex will appear in two differentfractions, and the identification of each fraction can be determinedaccording to the polarization measurement (i.e, the complex hassignificantly higher polarization than the free probe). It is alsopossible to run the free probe alone by the same separation method as anindicator to determine which fraction contains the free probe. Once theposition of the free probe is known, the complex can simply be detectedand quantitated by fluorescence measurement, such as laser-inducedfluorescence (LIF), without polarization.

[0104] It is preferable to separate the free probe from the complex inthe present invention. One reason is that the sensitivity with which todetect the complex is much higher if the method contains a separationstep. When the entire incubation mixture is subjected to polarization,the signal from the free probe is mixed with that from the complex, andthe net increase in polarization may not be very conspicuous. This isparticularly a problem if the probe is relatively large, resulting in asubstantial background polarization. If the free probe is separated fromthe complex, the background noise arising from the free probe iseliminated, and polarization due to complex formation can be detectedwith higher sensitivity.

[0105] A common problem with separating binding complexes is that weakbinding complexes tend to dissociate during separation. For example, inthe mobility shift method of detecting binding complexes, an incubationmixture is separated by gel electrophoresis, and the position of thecomplex is then located by detecting the label contained in the probe.However, due to the pH, temperature or electronic field to which acomplex is exposed during electrophoresis, a weak complex may dissociateduring electrophoresis. If so, the location of the probe would bemistakenly interpreted as the location of the complex.

[0106] This problem is not a concern in the present invention. Since thefluorescence polarization measurement is very different between acomplex and a free probe, it can be determined with significantcertainty whether a fraction contains a complex or a free probe.Furthermore, the present invention can be used to monitor complexformation on-line during the course of the separation. Therefore, thepresence of a complex can be detected before it dissociates.

[0107] A particular interesting application of the present invention islibrary screening. It is common to screen a large number of chemicalcompound libraries for a binding factor. Since the present inventionenables one to quickly determine if a binding factor exists in a libraryor not, without having to separate the free probe and the complex, it isvery useful in the initial screening. Once it has been determined whichlibraries contain binding factors, the incubation mixtures containingthe libraries of interest can be separated using an appropriate method,and the present invention can again be used to locate the fractionshaving the complex, thereby allowing identification of the bindingfactors.

[0108] Another particular application of the present invention is todetect nucleic acid damages. Nucleic acid damages, such as mismatches,breaks or the formation of DNA adducts, often occur after the nucleicacid is exposed to carcinogens. The present invention can be used todetect nucleic acid damages using a factor which is capable of bindingdamaged nucleic acids. Thus, a sample suspected of having damagednucleic acids can be incubated with a binding factor as well as afluorescently labeled oligonucleotide probe harboring the specificnucleic acid damage. The binding complex between the probe and thebinding factor can be detected using the present invention, with orwithout separation. If the sample contains the damage of interest, itwould compete with the probe for the binding factor to result in adecrease in binding complex formation between the probe and the factor.

[0109] Any factor capable of binding specifically to a damaged nucleicacid can be used in the present invention, such as any antibody whichrecognizes a specific DNA adduct; the UvrA and UvrB proteins which bindUV dimers, polycyclic aromatic hydrocarbon adducts, cis-platinumadducts, aflatoxin adducts, psoralen adducts, anthramycin adducts,mitomycin C adducts, N-acetoxy-2-aminofluorene adducts, andN-hydroxy-2-aminofluorene adducts; the DNA-dependent protein kinasewhich specifically bind to DNA double strand ends; poly(ADP-ribose)polymerase which binds to single-strand breaks and double-strand breaks;and the MutS protein which binds to several different mismatches (see,e.g. U.S. Pat. No. 6,132,968).

[0110] The following examples are offered to illustrate this inventionand are not to be construed in any way as limiting the scope of thepresent invention.

EXAMPLES

[0111] In the examples below, the following abbreviations have thefollowing meanings. Abbreviations not defined have their generallyaccepted meanings.

[0112] °C.=degree Celsius

[0113] hr=hour

[0114] min=minute

[0115] μM=micromolar

[0116] mm=millimolar

[0117] M=molar

[0118] ml=milliliter

[0119] μl=microliter

[0120] mg=milligram

[0121] μg=microgram

[0122] PAGE=polyacrylamide gel electrophoresis

[0123] rpm=revolutions per minute

[0124] FBS=fetal bovine serum

[0125] DTT=dithiothrietol

[0126] PBS=phosphate buffered saline

[0127] CE=capillary electrophoresis

[0128] LIFP=laser induced fluorescence polarization

[0129] LIF=laser induced fluorescence

[0130] PMT=photomultiplier tube

[0131] SSB=single-stranded DNA binding protein

[0132] FITC=Fluorescein isothiocyanate

[0133] SEA=staphylococcal enterotoxin A

[0134] FPIA=fluorescence polarization immunoassay

[0135] IAF=5-iodoacetamidofluorescein

[0136] RT=reverse transcriptase

[0137] TBE=Tris-borate-EDTA

[0138] FAM=carboxyfluorescein

[0139] AMV=avian myeloblastosis virus

[0140] MMLV=Moloney murine leukemia virus

[0141] BPDE=benzo[a]pyrene diol epoxide

[0142] TMR=tetramethylrhodamine

[0143] EOF=electroosmatic flow

[0144] LED=light-emitting diode

[0145] DMEM=Dulbecco's modified Eagle's medium

[0146] DMSO=dimethylsulfoxide

Section A: Examples 1-6 Application of the Present Invention in ComplexFormation Between Various Biological Molecules

[0147] Instrumentation. A laboratory-built capillary electrophoresiswith laser induced fluorescence polarization (CE/LIFP) detection systemwas used (Ye, et al., 1998). Electrophoresis was performed using a highvoltage power supply (Model CZE 1000R, Spellman, Plainview, N.Y.) and afused silica capillary (Polymicro Technologies, Phoenix, Ariz.). Thedetection end of the capillary was inserted in a sheath flow cuvette(NSG Precision Cells, Farmingdale, N.Y.). A plane polarized laser beamfrom an argon ion laser (Model 2014-65ML, Uniphase, San Jose, Calif.)was filtered through a laser line filter (488 nm, 10-nm band width,Newport, Fountain Valley, Calif.) and was used for excitation. The lightemitted during fluorescence was collected with a 60× microscopeobjective lens (0.7 NA, Universe Kogaku, Oyster Bay, N.Y.), filteredwith a narrow bandpass filter (515 nm, 10-nm band width, Newport), andpassed through a pinhole. The emitted light was subsequently split witha broadband polarizing beamsplitter cube (Melles Griot, Irvine, Calif.)into vertically and horizontally polarized components, which weredetected with two photomultiplier tubes (PMT1 and PMT2, R1477,Hamamatsu, Japan). The operation of the power supply and the acquisitionof data were controlled by a Power Macintosh computer with anapplication software written in LabView (National Instruments, Austin,Tex.).

[0148] There is no fundamental difference between this apparatus and theconventional fluorescence detectors that are also capable of anisotropymeasurements. The only difference is that our cuvette (flow cell) ismuch smaller (0.2×0.2 square) than commercially available cells and thatour detector is capable of handling the small volumes suitable for CEseparation.

[0149] CE/LIFP Analysis. Unless otherwise stated the CE/LIFP were run asfollows. A capillary of 20 mm i.d, 148 mm o.d. and 35 cm in length wasused in conjunction with 25 mM Na₂B₄O₇ (pH 9.1) as a typical runningbuffer. Prior to sample analysis, the capillary was preconditionedperiodically by successive rinsing with 0.1 M NaOH, deionized water andthe running buffer to ensure reproducibility of the separation. Sampleswere electrokinetically injected into the capillary by applying anelectric field of 143 V/cm for 5 s. Separation was carried out under anelectric field of 714 V/cm. The electrophoretic mobility (m) of a solutewas calculated using the following equation (Haugland, 1996; Schantz, etal., 1972).

m=L(1/t _(eo)−1/t)/E  (1)

[0150] where L and E are the capillary length and applied electric fieldstrength; t_(eo) and t are the migration times of the solvent andsolute, respectively.

[0151] Horizontally and vertically polarized fluorescence intensitiesmeasured by the LIFP detector were optimized by aligning a tightlyfocused laser beam with a small-diameter sample stream and by balancingsignals from the two PMTs. An aqueous solution of disodium fluorescein(10⁻⁹ M) was passed through a capillary inserted in a sheath flowcuvette. The sheath fluid, identical to the CE run buffer, wasintroduced into the cuvette hydrodynamically by keeping the inletreservoir of the sheath buffer 1 cm higher than the outlet reservoir.The vertically polarized laser beam was focused onto a spot about 20 mmbelow the tip of the capillary. The angle and position of the cuvetterelative to the detection optical path were adjusted so that roughlyequal signals with maximum outputs from both PMTs were achieved. Thevalues of fluorescence anisotropy (A) were calculated according to(Shimura, et al., 1994; Schultz, et al., 1993; Lakowicz 1983).

A=(I _(v) −I _(h)/(I _(v)+2I _(h))  (2)

[0152] where I_(v) and I_(h) are the fluorescence intensities ofvertically and horizontally polarized components, respectively.

[0153] The values of fluorescence polarization, P, were calculatedaccording to (Lakowicz 1983; Perrin 1926; Weber, 1953; Dandliker, et al.1970)

P=(I _(v) −GI _(h))/(I_(v) +GI _(h))  (3)

[0154] where I_(v) and I_(h) are the fluorescence intensities of thevertically and horizontally polarized components, respectively; G is anempirical constant that corrects for the polarization bias introduced bythe optics and the detection system. The G value was determined as theintensity ratio of vertical to horizontal polarization components offluorescein. For a well-balanced system, the polarization bias isrelatively small (G=0.98−1.0) and therefore, is negligible.

[0155] It is possible to have unequal transmission of the two orthogonalpolarizations through the emission optical trains and, therefore,unequal sensitivity of the PMT detectors for vertical and horizontalpolarized emission. To correct for this potential bias, the PMT voltagewas adjusted until the fluorescence intensities from the two PMT's (thevertically and horizontally polarized fluorescence) were identical fordilute fluorescein (10⁻⁹ M), which is assumed to have negligibleanisotropy.

[0156] In the first set of examples, laser induced fluorescencepolarization (LIFP) detector was used in conjunction with capillaryelectrophoresis to demonstrate the utility of CE/LIFP. Changes in theelectrophoretic mobility and fluorescence polarization of thefluorescent probe upon complex formation with the binding partner weremeasured simultaneously, thereby providing complementary information onthe binding interaction. This information could not be obtained witheither CE or LIFP used alone. Unless otherwise noted, the smallermolecule of a binding pair was labeled with a fluorophore such asfluorescein. The complex was formed by mixing the fluorescent substratewith the corresponding binding partner and was electrophoreticallyseparated from the unbound substrate followed by on-line detection withLIFP. Our results showed expected increases in fluorescence polarizationupon complex formation, demonstrating the usefulness of the technique inbinding studies involving a wide variety of biomolecules.

[0157] For binding systems that have low affinity, dissociation may takeplace during separation. We overcame this problem by including thebinding reagents in the CE separation buffer to stabilize the complex asdemonstrated in system in DNA-protein binding studies. A fluorescentlylabeled oligonucleotide or SSB protein was used as a probe and thebinding interactions with its partner were studied in either of twoformats, depending on the stability of the complexes formed. For weakbinding interactions, the binding partner was included in running bufferto stabilize the complexes during CE separation. The electrophoreticmobility and fluorescence anisotropy of the fluorescent probe weremeasured as a function of the concentration of its binding partner inthe running buffer. Both the electrophoretic mobility and fluorescenceanisotropy were used to determine the binding constants andcooperativity. For high affinity interactions, mixtures containing thefluorescent probe and its binding partner at varying ratios wereincubated prior to separation. The complexes formed off-column were thenseparated by CE with a running buffer free of the binding components.The electrophoretic mobility and fluorescence anisotropy measurementswere used for the identification of the complexes and for the study ofbinding stoichiometry.

[0158] Materials and Reagents. Disodium fluorescein of purified gradewas obtained from Fisher Scientific (Fair Lawn, N.J.) and was used forinstrument alignment. Fluorescein isothiocyanate (FITC) isomer I,L-tryptophan, staphylococcal enterotoxin A (SEA), polyclonal (rabbit)antibody to SEA, SSB protein, d(pT)₁₈, single-stranded M13mp8 phage DNAand FPIA (fluorescence polarization immunoassay) dilution buffer (pH7.4) containing phosphate, bovine protein and sodium azide, wereobtained from Sigma (St. Louis, Mo.).

[0159] Fluorescein-dUTP was obtained from Molecular Probes (Eugene,Ore.). A 5′-oligolabeling kit containing T4 polynucleotide kinase, ATPSand 5-iodoacetamidofluorescein (IAF) was obtained from AmershamPharmacia Biotech (Buckinghamshire, England).

[0160] Fluorescein labeled oligonucleotides 11-mer (5′-CGCGATACGCC-3′;SEQ ID NO:1) and 37-mer (5′-CCTTAAGCTTCCTCAACCACTTACCATACTCGAGATT-3′;SEQ ID NO:2) were provided by J. Lee of Cross Cancer Institute and T.Carnelley of Department of Public Health Sciences, University ofAlberta.

[0161] The fluorescein labeled vancomycin and polyclonal (sheep)antibody to vancomycin were from a Sigma diagnostics reagent set. Theactual compositions and concentrations of these solutions were notavailable. The trp repressor protein, trp operator DNA and trp bindingbuffer (pH 7.6) were obtained from PanVera (Madison, Wis.) as a tiprepressor-DNA binding kit. The trp operator was a 5′-fluoresceinlabeled, 25 base pair, oligonucleotide with sequence:

[0162] (5′-ATCGAACTAGTTAACTAGTACGCAA-3′)

[0163] (3′-TAGCTTGATCAATTGATCATGCGTT-5′)

[0164] Fluorescent Labeling of SEA. SEA was labeled with FITC and theextent of modification was estimated according to the methods describedby Brinkley, et al., 1992. A 10-fold molar excess of FITC was added to asolution of SEA (0.1 mg/mL) in 25 mM Na₂B₄O₇ (pH 9.1). The reaction wasallowed to proceed for 1 h at room temperature and then terminated byadding excess of hydroxylamine. The fluorescently labeled SEA wastransferred into a disposable dialyzer tube (Spectra/Por CE SterileDispoDialyzer, molecular weight cut-off 10,000 Da) and purified bydialysis against 10 mM sodium phosphate buffer (pH 7.4) at 4° C. for 2days. The degree of fluorescent labeling was determined by analyzing thefluorescently labeled SEA and the free dye in 25 mM Na₂B₄O₇ (pH 9.1)using a Hewlett-Packard (Palo Alto, Calif.) Model 1040A diode arraydetector. A Gilson (Villies le Bel, France) Model 307 HPLC pump was usedto introduce the sample solution. The molar ratio (R) of the fluorophoreto SEA protein was calculated according to the following equation:

R=A _(490, p) e _(p) /[A _(277, p) −A _(490, p)(A _(277, d) /A_(490, d))]e _(d)  (4)

[0165] where A_(277, p) and A_(490, p) are the absorbance offluorescently labeled SEA protein at 277 and 490 nm; A_(277, d) andA_(490, d) are the absorbance of the free dye at 270 and 490 nm; e_(p)and e_(d) are the extinction coefficients of SEA at 277 nm and the freedye at 490 nm, respectively.

[0166] Fluorescent Labeling of SSB protein and d(pT)₁₈. The SSB proteinwas labeled with FITC and the extent of modification was estimatedaccording to the methods described by Brinkley, et al., 1992. A 10-foldmolar excess of the dye was added to a solution of the SSB protein (1.4μg/mL) in 25 mM Na₂B₄O₇ (pH 9.1). The reaction was allowed to proceedfor 1 h at room temperature and then stopped by adding excess ofhydroxylamine. The FITC-labeled protein was purified using a prepackedbio-spin column (Bio-Gel P-6, Bio-Rad, Hercules, Calif.). The absorbanceof FITC (280 nm) and FITC-SSB (490 nm) in 25 mM Na₂B₄O₇ (pH 9.1) wasmeasured using a Hewlett-Packard (Palo Alto, Calif.) Model 1040A diodearray detector equipped with a Gilson (Villies le Bel, France) Model 307HPLC pump. The molar ratio of the fluorophore to protein was calculatedfrom the absorbance measurements using the following extinctioncoefficients: FITC (Fey, et al., 1984), e₄₉₀=73000 cm⁻¹ M⁻¹; and SSB(Thompson, et al., 1986), e₂₈₀=120000 cm ⁻¹ M⁻¹. Absorbance of FITC at280 nm was corrected for in the calculation of the molar ratio of FITCto protein. The labeling of d(pT)₁₈ at the 5′-end with5-iodoacetamidofluorescein was accomplished by following a protocolprovided by Amersham.

[0167] Formation of the Complexes. Various volumes (0, 2, 4, 6, 8 mL) ofantibody solution from a test kit for vancomycin were mixed with 10 mLaliquots of fluorescein-labeled vancomycin solution in 0.5 mLmicrocentrifuge tubes. FPIA dilution buffer was added to each tube to afinal volume to 200 mL. The tubes were vortexed for 30 s and the mixturewas allowed to incubate at room temperature for 15 min. Binding studiesfor SEA-antibody and trp repressor-operator systems were conductedsimilarly, with appropriate amounts of the binding partners. The sampleswere analyzed by CE/LIFP.

[0168] For weak interactions, protein-DNA complexes were formedon-column with excess amounts of the protein. Buffer solutionscontaining various concentrations of the SSB protein or oligonucleotidewere used as CE running buffers. The fluorescently labeled DNA wasinjected into the capillary for CE/LIFP analysis. For stronginteractions, protein-DNA complexes were formed off-column. Variousvolumes (0, 2.5, 5.0, 10, 15 mL) of the M13 phage DNA solution ( 43 nM)were mixed with 1.0-mL aliquots of FITC-SSB protein solution (12.5 mM)in 0.5-mL microcentrifuge tubes. FPIA dilution buffer was added to eachtube to a final volume of 50 mL. The tubes were vortexed for 30 s andthe mixtures incubated at room temperature for at least 15 min prior toCE/LIFP analysis.

Example 1

[0169] Peptide-Protein Interaction. Binding of vancomycin to itsantibody was chosen as an example because of the therapeutic importanceof vancomycin and because of the availability of its antibody as anaffinity agent. Vancomycin is a water soluble, tricyclic glycopeptideand is strongly bound to its antibody in solution as has beendemonstrated in a homogenous immunoassay (Schenzer, et al., 1983).

[0170] The complex, formed as described above, and the unboundvancomycin can be resolved by CE and can be readily identified based ontheir differential fluorescence polarization values. Twoelectropherograms were obtained from a single CE separation of a samplecontaining fluorescein-labeled vancomycin and anti-vancomycinantibodyand are shown in FIG. 1. Both vertically (I_(v)) andhorizontally (I_(h)) polarized fluorescence components were measuredsimultaneously. Fluorescein was added as a reference compound to correctfor any possible polarization bias of the instrument. Thefluorescein-labeled vancomycin and the fluorescein dye rotate rapidly insolution and exhibit little fluorescence polarization. Thus, thefluorescence intensities corresponding to the two polarized componentsare nearly equal. The binding of vancomycin to its antibody results in asubstantial increase in the molecular size and a slower rotation of themolecule. The complex exhibits significant fluorescence polarization.The intensity of the vertically polarized fluorescence (I_(v)) wassignificantly higher than that of the horizontal component (I_(h)) forthe complex. The same trend was observed with the complex formed atvarious vancomycin to antibody ratios. A mean fluorescence polarizationwas found to be 0.28±0.02. This value represents the intrinsicpolarization of the complex, which depends on rotational diffusion ofthe molecule but is independent of the amounts of the drug and antibodyadded.

[0171] The increase of fluorescence polarization upon complex formationcan be expected from the fluorescence polarization principle (Lakowicz1983; Perrin 1926; Weber, 1953; Dandliker, et al. 1970). A fluorescentmolecule, when excited by a polarized light, emits fluorescence with itspolarization (P) controlled by rotational correlation time (f) andfluorescence lifetime (t) as shown by the Perrin equation (Perrin, 1926)

(1/P−1/3)=(1/P ₀−1/3)(1+t/f)  (5)

[0172] where P₀ is the intrinsic polarization in the absence ofrotational diffusion. When the rotational correlation time is smallrelative to the fluorescence lifetime, the fluorescence is depolarized.When the fluorescence lifetime is constant for a given fluorophore(e.g., 4 ns for fluorescein), an increase in polarization may beobserved with increasing rotational correlation time. The rotationalcorrelation time can be estimated according to the Debye-Stokes-Einsteinequation (Gottfried, et al., 1999)

f=Mh(v+h)/RT  (6)

[0173] where M is the mass of the molecule, h is the viscosity of thesolution, v is the specific volume of the molecule, h is the degree ofhydration, T is the absolute temperature, and R is the ideal gasconstant. Using a typical specific volume (v) of 0.735 cm³/g and atypical value of hydration (h=0.2 cm³/g) (Gottfried, et al., 1999), weestimated the rotational correlation time of vancomycin (0.7 ns) and itscomplex with the antibody (58 ns). The reduced rotational diffusion ofvancomycin when bound to the antibody resulted in an increase influorescence polarization from 0.08 for the free vancomycin (1838 Da) to0.28 for the antibody-bound vancomycin (˜152,000 Da). The molecularweight, the estimated rotational correlation time and the observedfluorescence polarization are summarized in Table 1. TABLE 1 Molecularweight, estimated rotational correlation time and observed fluorescencepolarization of FITC labeled substrates and their protein complexesEstimated Molecular Rotational Observed Weight Correlation TimeFluorescence Substrate and Complex (Da) (ns)^(a) Polarization FITClabeled substrates Vancomycin 1,838 0.7 0.08 ± 0.02 Trp Operator 15,0005.8 0.11 ± 0.02 SEA 28,000 11 0.14 ± 0.02 Complexes Vancomycin-Antibody152,000 58 0.28 ± 0.02 Trp Operator-Repressor 30,000 12 0.25 ± 0.02SEA-Antibody 180,000 69 0.14 ± 0.02

[0174] As intrinsic property of a molecule, the characteristicfluorescence polarization provides evidence for the binding of thesubstrate to its antibody without the need of tedious titrationprocedures, thereby promising considerable savings on time and reagents.This feature makes the CE/LIFP approach particularly suitable forapplications such as screening specific monoclonal antibodies where thespeed and convenience are the major concerns when choosing a screeningmethod (Barret, 1994).

Example 2

[0175] Protein-Protein Interaction. Binding of SEA to its antibody wasstudied with an intention of developing CE based immunoassays for thisnatural toxin. SEA has been known for many years to cause food poisoning(Marrack, et al., 1990) and, therefore, it is of considerable publichealth interest to develop rapid and sensitive analytical methods.Radioimmunoassays and enzyme-linked immunosorbent assays for this toxinhave been described in the literature (Johnson, et al., 1973; Fey, etal., 1984; Thompson, et al., 1986).

[0176] Experiments were run to characterize the FITC labeled SEA and toexamine the formation and stability of its complex with thecorresponding antibody. Using equation (1) and literature values ofextinction coefficients for the dye (e_(d)=73,000 cm⁻¹ M⁻¹) and theprotein (e_(p)=40,900 cm⁻¹ M⁻¹),(Haugland, 1996; Schantz, et al., 1972)we estimated the molar ratio of the dye to the protein to be 5.9±0.3 forthe labeled SEA.

[0177] Fluorescently labeled SEA exhibits an appreciable polarization(0.14), making it readily identifiable even in the presence of theresidual dyes. This finding suggests that the CE/LIFP may be used tomonitor the progress of labeling reactions and to verify the purity ofthe reaction products. The broadness of the peak is attributed to theheterogeneity of the protein (Schantz, et al., 1972) as well as to thepresence of multiply labeled products (Craig, et al., 1998).

[0178] The electropherograms from a CE/LIFP analysis of a mixture ofFITC-SEA and its antibody show a new peak attributable to the complexbetween fluorescently labeled SEA and its antibody. The complexexhibited measurable fluorescence polarization (0.14) as seen with thefree SEA. However, there was no net increase of polarization uponcomplex formation. This is not surprising given the relatively highmolecular weight of SEA itself (28,000 Da). In an aqueous solution atroom temperature, the rotational correlation time (f) of SEA isestimated using equation (6) to be approximately 11 ns, which is about2.5 lifetimes of fluorescein (t, less than 4 ns). With such a longrotational correlation time, the t/f term (<0.4) in equation (5)contributes little to the observed fluorescence polarization (P). Thepolarization may approach to P₀, the intrinsic value for the moleculebecause it is known that fluorescence polarization generally approachessaturation with molecular weight beyond 20,000 Da (Guo, et al., 1998).As a result, further increase in correlation time due to the binding ofantibody did not lead to any significant increase in polarization. Thepolarization of 0.14 for the complex is well below the theoretical limitof 0.5, indicating that local rotation of the fluorophore may occurwithin the SEA molecule.

[0179] Because of no increase in polarization noted in this case,additional evidence is needed to confirm the complex formation, whichcan be obtained by titrating a fixed amount of FITC-SEA with varyingamounts of the antibody. As expected, the complex peak increases at theexpense of the unbound SEA peak (Wan, et al., 1999)

Example 3

[0180] Protein-DNA interaction. The interaction between trp operator(DNA) and trp repressor protein of Escherichia coli serves as awell-characterized system for gene expression and regulation (Crawford,et al., 1980). In the presence of tryptophan, the repressor proteinbinds with high affinity to the operator sequence found within thepromoter region of the trpEDCBA operon and represses transcription ofthose genes whose protein products are responsible for the synthesis oftryptophan. In the absence of tryptophan, trp repressor is inactive andthe trp operon is expressed, resulting in the biosynthesis oftryptophan. The binding of the trp operator to trp repressor has beenstudied extensively (Carey, 1988; Otwinowski, et al., 1988;Lawson, etal., 1988; LeTilly, et al., 1993; Zhang, et al., 1994; Stebbins, et al.,1996).

[0181] Two formats were used to explore the potential of CE-LIFP in thestudy of DNA-protein interactions in the trp operator (DNA) and trprepressor protein system. In the first format, the affinity complex wasformed dynamically in the separation capillary and maintained atequilibrium with the free protein during the electrophoresis. In thesecond format, the complex was preformed by incubation and thenseparated from the unbound protein molecule.

[0182] The electropherograms of fluorescently labeled trp operatoroligonucleotide obtained with and without the trp repressor protein inthe running buffer were compared. The I_(v), and I_(h), representingvertically and horizontally polarized fluorescence, respectively weremeasured. In the presence of the trp repressor, a tailed peak withmigration time of 4.6 min was observed. This peak corresponds to thecomplex formed between the trp operator and trp repressor judging fromthe fluorescence polarization that increased from 0.11 (without the trprepressor in the running buffer) to 0.25 (with the inclusion of the trprepressor in the running buffer).

[0183] It is noted that fluorescently labeled DNA displays multiplepeaks in both cases and that these peaks do not disappear even in thepresence of excess trp repressor. The persistence of the multiple peakssuggests the presence of multiple DNA structures (Stebbins, M. A , etal., 1996; Hamdan, et al., 1998), some of which are not recognized bythe repressor protein. Experiments have established the dynamicformation of the DNA-protein complex (Wan, et al., 1999).

Example 4

[0184] SSB protein/ssDNA. The SSB protein plays an important role in theDNA replication, recombination and repair proces (Bandyopadhyay, et al.,1978; Krauss, G., et al., 1981; Chase, et al., 1986; Lohman, et al.,1994) although mechanisms for its functions in these processes have notyet been elucidated. The SSB protein exists as a tetramer in solutionwith a subunit weight of about 20000. It binds cooperatively tosingle-stranded DNA (approximately 32-60 nucleotides per protein),keeping the DNA in an extended configuration and protecting it fromnuclease digestion. The results shown in the SSB/DNA binding experimentsdemonstrate the resolving power and identification capabilities ofCE/LIFP in these systems.

[0185] CE/LIFP may be used in binding studies through the measurement ofchanges of either electrophoretic mobility or fluorescence anisotropy ofa binding component upon affinity interactions. Usually, the bindingcomponent of lower molecular weight is used as a probe so as to inducegreater mobility or anisotropy changes upon binding. In this example,both fluorescently labeled DNA fragments and binding protein were usedas probes for the study of protein-DNA interactions.

[0186] In this example, a fluorescein labeled 11-mer (F-11-mer) wasevaluated as a probe for examining its binding with the SSB protein.Electropherograms of F-11-mer in the absence and presence of the SSBprotein in the running buffer were run and the vertical (I_(v)) andhorizontal (I_(h)) components of fluorescence were measuredsimultaneously for each. Fluorescein labeled dUTP (F-dUTP) was used as areference compound to correct for possible fluctuations inelectroosmotic flow and unequal detection sensitivity between the twodetection channels of the instrument. In the absence of the bindingprotein, the fluorescent probe has an electrophoretic mobility ofm=3.99×10⁻⁴ cm² V⁻¹ s⁻¹ and a fluorescence anisotropy of A=0.05. In thepresence of 0.7 mM of the SSB protein in the running buffer, theelectrophoretic mobility of the oligonucleotide probe is reduced to1.93×10⁻⁴ cm² V⁻¹ s⁻¹ whereas the anisotropy is increased to 0.25. Thedecrease in electrophoretic mobility and increase in anisotropy are dueto the binding of the F-11-mer with the SSB protein. In contrast, themobility and anisotropy of F-dUTP are essentially unchanged, consistentwith the fact that the SSB protein has very low binding affinity for themononucleotide.

[0187] The electrophoretic mobility and anisotropy changes observed inthe experiments arise from the effects of the binding protein on themolecular motion of the probe. In the absence of the binding protein,the fluorescently labeled oligonucleotide probe migrates with a mobilitysimilar to that of F-dUTP in the free zone electrophoresis mode. It hasa low fluorescence anisotropy because of its small molecular size (MW4000) and random motion in solution. When bound to the SSB protein (MW80000), the size of the fluorescent molecule is markedly increased,resulting in a slower molecular motion in the solution. Therefore, it isnot surprising that binding of the SSB protein to the oligonucleotideprobe gives rise to a marked increase in anisotropy. The mobility andanisotropy of the complex approach to those of the binding protein(m=1.07×10⁻⁴ cm² V⁻¹ s⁻¹ and A=0.23 for FITC-SSB. While theelectrophoretic mobility of a compound is proportional to its charge tomass ratio, (Chu, et al., 1995; Baker, 1995) the fluorescence anisotropyis mainly determined by its molecular size, shape, and fluorescencelifetime (Lakowicz, 1999).

[0188] The mobility and anisotropy of the F-11-mer were measured withthe running buffer containing various concentrations of the SSB protein.Nanoliter amounts of the F-11-mer (10⁻⁹ M) were injected into thecapillary that was filled with the running buffer containing 10⁻⁷-10⁶ MSSB protein. Thus, the concentration of the binding protein in therunning buffer was in large excess and was not affected significantly byits binding with the F-11-mer. Under this condition, the quantitativeinterpretation of the binding profiles can be carried out using astandard four-parameter logistic equation (Motulsky, 1999):

y=(a+bK ^(n) x ^(n))/(1+K ^(n) x ^(n))  (7)

[0189] where y is the observed response such as electrophoretic mobilityor fluorescence anisotropy of the oligonucleotide probe at a givenconcentration of the binding protein, x; K is the apparent bindingconstant; a and b are the responses of the free and bound probe,respectively; and superscript n is the Hill coefficient describing thesteepness of the curve. The experimental data were fitted to the aboveequation using nonlinear regression analysis (SigmaPlot, version 4, SPSSInc.). Binding constant (K) measurements based on mobility andanisotropy of the F-11-mer were similar, which were approximately4.4×10⁶ M⁻¹ and 5×10⁶ M⁻¹, respectively. These values are comparable toprevious measurements by other methods. For example, Molineux (Molineux,et al, 1975) reported that SSB has an affinity of about 2×10⁶ M⁻¹ ford(pT)₈ and Krauss (Krauss, et al, 1981) reported an affinity of 1.4×10⁶M⁻¹ for d(pT)₁₆ using fluorescence quenching methods. Binding constantsand fitting parameters from nonlinear regression analysis are summarizedin Table 1.

[0190] Furthermore, binding interactions involving protein-DNA complexesthat differ in stoichiometry can be accomplished with thisinstrumentation. Fluorescein labeled 37-mer (F-37-mer) was chosen as aDNA probe since its complexes with the SSB protein are of higherstability, allowing examination of a distribution of different speciesin the binding interactions. Electropherograms of F-37-mer with theabsence and presence of the SSB protein in the CE running buffer werecollected. Changes in electrophoretic mobility and fluorescenceanisotropy upon formation of complexes between F-37-mer and SSB proteinare observed as expected. It is noted that the initial single peak ofF-37-mer was split into two when bound to the SSB protein. Both complexpeaks display strong fluorescence isotropy (A=0.23). These are likely2:1 (peak 2) and 1:1 (peak 1) protein-DNA complexes. For the 1:1 bindinginteraction, variations of the m and A values for the F-37-mer with thebinding protein concentration were compared, from which the apparentbinding constants were obtained. Again, both mobility and anisotropymeasurements gave very similar results, with binding constants ofapproximately 2×10⁷ M⁻¹ (see Table 1). This is approximately 5-foldincrease in binding affinity of SSB for the 37-mer compared to itsbinding with the 11-mer. This is consistent with the contribution ofcooperativity to the binding strength. The SSB protein is a tetramer andthe number of the binding sites of the SSB protein varies with thelength of the oligonucleotides because each of the four subunits of theprotein covers 6-8 nucleotides (Krauss, et al, 1981; Chase, et al.,1986). While one subunit may bind to the 11-mer, all the four subunitsof the SSB tetramer could bind to the 37-mer, resulting in thecorresponding increase in binding constant. Because the two complexspecies were not well resolved particularly at low proteinconcentrations, there was a relatively large uncertainty associated withmobility and anisotropy measurements for the 2:1 complex. Consequently,we were unable to precisely determine the corresponding binding constantfor the 2:1 complex.

Example 5

[0191] Fluorescein Labeled DNA Binding Protein as a Probe. Fluorescentlylabeled DNA oligonucleotides have exclusively been used as probes in theanalysis of protein-DNA interactions by gel retardation, CE or FPtechniques. However, many bioanalytical applications require the use ofa fluorescently labeled binding protein for its ability to recognize andbind to specific structures of DNA. In this example, FITC-labeled SSBprotein interacts with a synthetic oligonucleotide and a single-strandDNA. The FITC-labeled SSB protein was prepared as described in theexperimental section and the dye to protein molar ratio was 5.3. In asolution of 25 mM disodium tetraborate with pH 9.1, the labeled SSBprotein displayed an electrophoretic mobility of 1.07×10⁻⁴ cm² V⁻¹ s⁻¹and a fluorescence anisotropy of 0.23.

[0192] Electropherograms of FITC-SSB protein were obtained with runningbuffers containing varying amounts of oligonucleotide, d(pT)₁₈. Bothvertically and horizontally polarized fluorescence emissions wereacquired simultaneously. There was only a slight increase influorescence anisotropy (DA>>0.02) of the protein probe upon binding tothe oligonucleotide. This is in accordance with the fact thatfluorescence anisotropy of a probe generally approaches saturation whenthe probes molecular weight exceeds 20000 (Lakowicz, 1999; Wan, et al.,1999; Guo, et al., 1998).

[0193] It is noted that formation of the complex gives rise to somesignificant changes in the mobility and peak shape of the FITC-labeledSSB protein. As the concentration of d(pT)₁₈ in the running bufferincreases, the electrophoretic mobility of the SSB protein increaseswith the peak becoming increasingly dispersed. The mobility increase ofthe low mobility species (the FITC-SSB protein, m=1.07×10⁻⁴ cm² V⁻¹ s⁻¹)is due to its binding to a high mobility DNA (m=2.19×10⁻⁴ cm² V⁻¹ s⁻¹for fluorescein labeled d(pT)₁₈). The peak broadening suggests that theformation of multiple protein-DNA complexes is possible in the presenceof increasing amount of DNA. A single protein molecule may bind severalDNA molecules in the presence of excess DNA. The complexes of varyingprotein to DNA ratios co-migrate in the separation capillary as a broadband. To clarify this point, we chose a DNA fragment much longer thand(pT)₁₈ to form stable complexes with the SSB protein. Because ofincreased stability, the multiple complexes formed off-column can beseparated without the need for adding the binding partner to the runningbuffer.

Example 6

[0194] Detection of labeled protein binding to ssDNA. In the followingembodiment, single-stranded M13mp8 phage DNA (7229 bases) was selectedas a binding partner for the labeled SSB protein. Varying amounts of thephage DNA were incubated with a series of binding solutions containing afixed amount of the FITC-labeled SSB protein. The mixtures were thenanalyzed by CE/LIFP with a running buffer free of the bindingcomponents. Separations of FITC-SSB protein and its complexes with theDNA at various molar ratios of DNA to protein show that as the amount ofDNA in the reaction mixture increases, new peaks emerge and becomeincreasingly retarded and broadened. The broad and multiple peaks withincreasing migration times showed strong fluorescence anisotropy(A=0.25), indicating the presence of multiple protein-DNA complexes.Three major peaks with increasing mobilities (peak 2, 2.16×10⁻⁴ cm² V⁻¹s⁻¹; peak 3, 2.66×10⁻⁴ cm² V⁻¹ s⁻¹; and peak 4, 3.16×10⁻⁴ cm² V⁻¹ s⁻¹)corresponded to the complexes with increasing DNA to protein ratios.

[0195] In the absence of the DNA, peak 1 corresponds to the SSB proteinprobe which has a m=1.07×10⁻⁴ cm² V⁻¹ s⁻¹. Addition of the DNA to theprotein (with DNA-to-protein ratio of 0.008 and 0.016) causes a decreasein peak 1 and the appearance of peak 2 (m=2.16×10⁻⁴ cm² V⁻¹ s⁻¹ ),indicating the formation of DNA-protein complex. With further increaseof DNA-to-protein ratio, the mobilities of the complexes (2.66×10⁻⁴ cm²V⁻¹ s⁻¹ for peak 3; and 3.16×10⁻¹ cm² V⁻¹ s⁻¹ for peak 4) shift towardsthat of the DNA (m=3.50×10⁻⁴ cm² V⁻¹ s⁻¹).²⁷ Because the amount of theprotein was fixed in this series of experiments, increasing amounts ofDNA in the reaction mixture favor the formation of complexes ofincreasing DNA:protein ratio. This example demonstrates an applicationof CE/LIFP to study multiple-complexes.

Section B: Examples 7-10 Complex Formation Between HIV-RT and Aptamers

[0196] In the next few examples the detection of human immunodeficiencyvirus type 1 reverse transcriptase was accomplished using aptamers asprobes in affinity capillary electrophoresis and laser inducedfluorescence polarization. CE determination of HIV-1 RT using anoncompetitive affinity assay has several advantages in terms of vastlydecreasing analysis time and involves much simpler chemical procedures.A fluorescently-labeled aptamer such as RT 12 or RT 26 eliminates theneed for the use of radio-labeled materials, and provides the firstdirect assay for HIV-1 RT. Since the aptamers were evolved to bindselectively to HIV-1 RT, interferences from RTs of other species waseliminated or greatly attenuated. Used in conjunction with otherlaboratory procedures correlating HIV-1 RT activity to viral loads, theassay could prove useful in the determination of HIV-1 viral load.

[0197] Apparatus: The CE/LIF instrument used in this work is the same asdescribed above except that a 543.5 nm green He—Ne laser (Melles Griot,Irvine, Calif., USA) with a 5 mW maximum out put was used as theexcitation source.

[0198] Reagents: All solutions were prepared using 18.2 MW distilled,deionized water (DDW) from a Milli-Q Gradient 10 Water System(Millipore, Nepean, Canada). Tris-borate-EDTA (TBE) (0.089 M tris, 0.089M boric acid, 0.0025M EDTA, pH 8.3), tris-glycine (0.025 M tris, 0.192 Mglycine, pH 8.3) and disodium tetraborate buffers (0.1 M, pH 9.1) wereprepared using reagent-grade materials and diluted to desiredconcentrations with DDW prior to being filtered through a 0.22 μm filterto remove particulate matter. The RT 12 aptamer(5′-ATCTACTGGATTAGCGATACTCGATTAGGTCCCCTGCCGCTAAACCATACCGCGGTAACTTGAGCAAAATCACCACTGCAGGGG-3′;SEQ ID NO:3) and the RT 26 aptamer(5′-ATCCGCCTGATTAGCGATACTTACGTGAGCGTGCTGTCCCCTAAAGGTGATACGTCACTTGAGCAAAATCACCTGCAGGGG-3′;SEQ ID NO:4) were labeled with 5′-FAM (5′-carboxyfluorescein) at theUniversity Core DNA Services, University of Calgary, Canada. HIV-1 RTwas obtained from Worthington Biochemicals (Lakewood, N.J.). RTs fromthe enhanced avian myeloblastosis virus (AMV) and the Moloney murineleukemia virus (MMLV) were obtained from Sigma (Mississauga, Canada).Cell culture media (RPMI with 10% fetal bovine serum (FBS)) was obtainedfrom the Cross Cancer Institute at the University of Alberta.

[0199] Capillary Electrophoresis/Laser-Induced Fluorescence: Uncoatedfused silica capillaries (20 μm I.D., 150 μm O.D.) were cut to a lengthof 40 cm and inserted into the sheath flow cuvette where the laser beamwas focused. Samples were injected for 5 s at a voltage of 15 kV (375V/cm), and electrophoresis was carried out at a running voltage of 20 kV(500 V/cm). The running buffer utilized for all experiments was 1×trisglycine. The laser power was set at 4 mW throughout. Periodically, thecapillaries were treated by running 0.1 M NaOH through the system at anrunning voltage of about 100 V/cm for 30 mintues, followed by therunning buffer (1×tris glycine) at 500 V/cm, to remove protein materialadsorbed on the capillary wall.

[0200] Affinity Complex Formation: The RT 12 aptamer and RT 26 aptamerwere received in 0.020 μg and 0.040 μg quantities, respectively. Thesewere diluted to 60 μL in 1×TBE in 600 μL microcentrifuge tubes andstored in a freezer at 20° C. when not in use, as were the RTs of HIV-1,AMV and MMLV. Stock solutions of 80 nM for the RT 12 aptamer and 170 nMfor the RT 26 aptamer were prepared in 1×TBE in 600 μL microcentrifugetubes. A 1000 nM stock solution of HIV-1 RT was similarly prepared inDDW. Complex formation was carried out in an incubation buffer of 1×TBE.The desired concentration of aptamer and protein was obtained bypipeting the appropriate volumes of aptamer and protein stock solutionsinto a 60 μL volume in 600 μL microcentrifuge tubes. The tubes were thenvortexed for 30 s and put on ice for about 5 minutes prior to sampleinjection into the capillary. All stock solutions were stored at 20° C.when not in use and all samples were kept on ice during the course ofexperimentation.

[0201] Interference Studies: To determine the degree to which theaptamers would bind with RTs from AMV and MMLV, experiments wereconducted in which complex formation experiments, as described above,were undertaken with AMV and MMLV RTs substituted for HIV-1 RT.Furthermore, complex formation experiments were conducted with RTs ofHIV-1, AMV and MMLV mixed together, with the AMV and MMLV RTs at thesame or higher concentration than HIV-1 RT. To determine the degree towhich matrix effects from cell culture media would interfere withcomplex formation, aliquots of RPMI containing 10% FBS were added tosamples containing both HIV-1 RT and aptamer, as well as aptamer alone.

Example 7

[0202] Detection of Affinity Complex Formation after CE Separation. Theaffinity complex was formed by adding increasing concentrations ofaptamer to a fixed concentration of HIV-1 RT (50 nM). In the absence ofHIV-1 RT, the aptamer peak is sharp with a migration time of around 4.2minutes. Tailing, a characteristic of the aptamer peak, was observedeven at the lowest aptamer concentrations used (1.7 nM). This mostlikely results from impurities in the DNA, as has been previouslyobserved (German, et al., 1998). These DNA impurities likely containmultiple DNA structures, which cannot be recognized by the HIV-1 RTprotein (Wan, et al., 1999; Stebbins, et al., 1996; Hamden, et a.,1998).

[0203] The RT-26-HIV-1 affinity complex was formed. The peakcorresponding to the complex had a migration time of about 3.3 minutesand was well resolved and Gaussian in appearance. The lack of featuressuch as bumps or shoulders suggested that the affinity complex wasprimarily of single stoichiometry. At concentrations of 50 nM HIV-1 RTand 17 nM RT 26, the complex peak migrated as a doublet or a shoulderappeared, indicating that RT 26 and HIV-1 RT were forming a complex oftwo stoichiometries. The peak area of the HIV-1 RT-aptamer affinitycomplex increases with increasing aptamer concentration.

[0204] Experiments were also undertaken using RT 12 at 8, 15, and 20 nMconcentrations added to a solution containing a fixed concentration of50 nM of HIV-1 RT. The results parallel those described above, in thatthe CE peak area of the affinity complex increased with increasingaptamer concentration. However, the RT 12-HIV-1 RT affinity complexexhibited two distinct peaks, one forming at the same migration time asthe RT 26-HIV-1 RT complex, while an additional early peak formed atabout 2.9 minutes. HIV-1 RT is a heterodimer of total molecular weight(120 kDa) with two sub-units of molecular weight 51 kDa and 66 kDa.Although it is possible that the dimeric forms of HIV-1 RT may bebinding to different sites on the differently-structured RT 12 aptamer,a more likely explanation is that the RT 12 is being incorporated intothe affinity complex in such a way as to produce a complex of twodifferent stoichiometries (Wan, et al., 1999). Later experiments, inwhich the RT 12-HIV-1 RT affinity complex was observed to migrate as asingle peak, confirm the belief that an affinity complex of differentstoichiometries was formed in these experiments. Because of the higherbinding constant of the RT 26 aptamer (81-mer), it was chosen for allfurther work.

Example 8

[0205] Use of CE/LIFP to create Calibration Curves: Calibration curvesfor HIV-1 RT were constructed using aptamer concentrations of 17 nM and60 nM, preferebly, 17 nM of the RT 26 aptamer. The RT 26 probe peak areadecreases with increasing HIV-1 RT concentration, reaching a limitingvalue at 100 nM and disappearing completely at 800 nM of HIV-1 RT. Thatthe aptamer was completely incorporated into the affinity complexindicates the aptamer was at its preferred orientation in the TBEincubation buffer, without the need for heat denaturing and the presenceof Mg²⁺ salts, as was found in another assay in which DNA aptamers wereused to bind IgE and thrombin (German, et al., 1998). Most samplesolutions were stable for about two weeks if immediately frozen afteruse, after which both the probe and affinity complex peak areas weresignificantly diminished. At HIV-1 RT concentrations of 7 nM or lower,it was necessary to perform experiments within 30-40 minutes becauseaptamer and complex peak area began to deteriorate. Because of practicalconsiderations such as these, calibration curve samples were preparedsequentially, and samples were prepared fresh daily.

[0206] The calibration curve for the bound complex showed an initialsteep increase in fluorescence intensity with HIV-1 RT concentration,followed by leveling off, indicative of binding saturation, beginning atabout 100 nM of HIV-1 RT. For the aptamer probe peak, this was mirroredby a similar steep loss in fluorescence intensity, followed by aleveling off at about 100 nM. It is incorporated in the affinitycomplex, ultimately completely, at 800 nM of HIV-1 RT. The steeplyrising sections of the curves were then investigated to determineanalytical utility. The linear dynamic range for both probe and complexpeaks extends to 50 nM of HIV-1 RT. In the case of the aptamer probe,least-squares linear regression provides a best-fit line having acorrelation coefficient (r²) of 0.985 and a slope of 0.612, whereas thesame fit to the bound complex provided an r² value of 0.986 and a slopeof −0.938. Relative standard deviations range from a high of 7.1% tomajority of 1.9%-2.5% for both probe and complex peaks.

Example 9

[0207] Use of CE/LIFP in Specificity determination: Experiments wereperformed in which all or some of the RTs were added together with HIV-1RT and 17 nM of RT 26 aptamer. AMV RT is present at a concentrationcomparable to that of HIV-1 RT, whereas, MMLV RT was an order ofmagnitude more concentrated and was increased to over two orders ofmagnitude above that of HIV-1 RT. The peak areas as measured by CE ofthe unbound aptamer did not decrease, remaining essentially identical tothat in the presence of HIV-1 RT alone. These results indicate that thepresence of other RT proteins, such as AMV-RT and MMLV-RT, does notaffect the determination of HIV-1 RT.

[0208] The RTs of AMV and MMLV can be shown not to cross-react with theaptamer and to be specific for HIV-1 RT. Using high concentrations of RT(4-4000 units/μL) and aptamer probe (140 nM), AMV-RT concentrations of 4units/μL, and MMLV-RT concentrations of 4000 units/μL, The peak area ofthe unbound aptamer as measured by CE is essentially the same in theabsence and presence of AMV-RT and MMLV-RT.

Example 10

[0209] Use of CE/LIFP to show Effects of Sample Matrix:Electropherograms from the analysis of mixtures containing 20 nM HIV-1RT and 17 nM RT 26 aptamer in TBE buffer, in RPMI cell culture mediumand in 100-fold dilute culture medium were measured. Undiluted culturemedium clearly affected the formation and CE/LIF analysis of thecomplex. This is not surprising because the RPMI culture medium wassupplemented with 10% FBS. This protein is known to affect HIV-1 RT(Lee, et al., 1987). When the cell culture medium was diluted 100-fold,the matrix interference on the complex formation and CE/LIF analysis wasminimal. The analysis of mixtures of the RT 26 aptamer and HIV-1 RT inTBE buffer and in 100-fold dilute culture media show similarelectropherograms.

Section C: Examples 11-21 Detection of Damaged DNA

[0210] In the next set of examples, the detection of DNA adducts ofbenzo[a]pyrene using immuno-electrophoresis with laser-inducedfluorescence is demonstrated on the analysis of A549 cells.Benzo[a]pyrene belongs to a class of compounds called Polycyclicaromatic hydrocarbons (PAHs), which are known exhibit strongcarcinogenic properties, presumably as a result of the damage that theyor their metobolites cause cause to DNA. In vivo, B[a]P is converted tobenzo[a]pyrene diol epoxide (BPDE). Because of the biologicalsignificance of DNA damage and repair, many techniques have beendeveloped for the determination of DNA damage (Pfeifer, 1996).

[0211] Synthetic BPDE-DNA adduct was used as a standard probe in acompetitive assay to determine the levels of BPDE-DNA adducts in a humanlung carcinoma cell line exposed to BPDE. A fluorescently labeledBPDE-DNA adduct standard and a BPDE-specific antibody were added to asample containing unknown amount of unlabeled BPDE-DNA adduct. Theunlabeled BPDE-DNA adduct and the labeled BPDE-DNA adduct compete toform complexes with the antibody. CE separation of the bound and unboundadducts allows determination of the bound concentration, which in turnis related to the amount of BPDE-DNA adduct in the sample. In contrastto other methods of performing immunoassays, CE-LIF allows rapidanalysis, excellent mass sensitivity and potential for automation. Thepopularity of this technique in immunoassays is well reflected innumerous reports, primarily for the determination of therapeutic drugs(Schulz, et al., 1995; Schmalzing, et al., 1995; Chen, et al., 1994;Evangelista, et al., 1994; Chiem, et al., 1998).

[0212] Preparation of BPDE-DNA adduct standard. BPDE powder(benzo[a]pyrene-r-7, t-8-dihydrodiol-t-9, 10-epoxide (+/−) (anti) wasobtained from Midwest Research Institute (Kansas City, Mo., USA. MRI0477; Lot CSL-98-775-17-16). The BPDE powder was dissolved in dimethylsulfoxide (DMSO) to a stock solution of 3 mM. A 16-mer oligonucleotide,5′-CCCATTATGCATAACC-3′ (SEQ ID NO:5), was treated with BPDE at a molarratio of 1:5 (oligonucleotide: BPDE), using a protocol similar to thatdescribed by Cosman (Cosman, et al., 1990). The oligonucleotide wasreconstituted in a buffer containing 20 mM phosphate/1.5% triethylamineat pH 11. To the oligonucleotide, a BPDE solution was added to a finalconcentration of 270 mM. The final mixture was incubated in the dark atambient temperature overnight with gentle shaking. Purification of theBPDE-modified oligonucleotide was carried out in two separate rounds ofHPLC elution using a preparative column (Phenomenex, Torrance, Calif.,USA. LUNA Su C18(2); 250×10 mm 5 mm particle size). In the first round,an isocratic elution using 70% methanol and 30% of 20 mM phosphate, pH 7was used to purify the BPDE-oligonucleotide by separating theoligonucleotides from the unreacted BPDE. The eluent containing theBPDE-oligonucleotide and the unmodified oligonucleotide was freeze driedand subjected to a second round of HPLC purification using a gradientelution of methanol/20 mM phosphate at pH 7. This purification stepseparates the BPDE-oligonucleotide from the unmodified oligonucleotide.The freshly purified BPDE-oligonucleotide was subjected to a standardkinase reaction to facilitate subsequent ligation to 5 otheroligonucleotides to form a BPDE-DNA duplex of 90 base pairs. TheBPDE-DNA duplex was gel purified using a 7.5% native polyacrylamide gel,and subsequently subjected to UV and fluorescence scanning to measureDNA concentration as well as to confirm the presence of BPDE moiety onthe 90-mer.

[0213] Specific monoclonal antibodies. Monoclonal antibodies 8E11 and5D11 were obtained from BD PharMingen (San Diego, Calif., USA). Bothantibodies were derived from BALB/c mice immunized with racemicanti-BPDE modified guanosine conjugated with bovine serum albumin(Santella, et al., 1984).

[0214] Preparation of BPDE-DNA adducts from A549 cells. A human lungcarcinoma cell line (A549) was incubated with BPDE to produce DNAadducts in genomic DNA. Briefly, the cell line was maintained inDMEM/F12 medium (Gibco BRL, Gaithersburg, Md., USA) supplemented with10% fetal bovine serum. The cells were seeded at 1×10⁵ cells per plateand maintained at 95% humidity and 5% CO₂ for 20 hours prior to theaddition of BPDE. Treatment of BPDE was carried out in duplicate sets ofA549 cells. Old culture media were removed from each culture plate andthe cells were washed twice with phosphate buffered saline (PBS). Mediacontaining BPDE at various concentrations (9.4, 18.8, 37.5, 75, 150, and300 mM final concentration) were added accordingly to the designatedplates. The cells were further incubated in the media containing BPDEfor 2 hours. The cells were then washed with PBS prior to the additionof DNAzol lysis reagent (Gibco BRL) to facilitate cell lysis. Subsequentsteps involved a standard 99.9% ice cold ethanol precipitation and a 70%cold ethanol wash to purify the genomic DNA. The final DNA pellet wasdissolved in distilled deionized water (ddH₂O) and DNA concentration wasmeasured at OD₂₆₀ using ddH₂O as a blank.

[0215] CE-LIF Instrumentation. The instrument for capillaryelectrophoresis with laser induced fluorescence detection is describedabove, with one modification. A 543.5 nm green He—Ne laser (MellesGriot, Irvine, Calif., USA) with a 5 mW maximum output was used as theexcitation source. In addition, in place of the polarizing beamsplitter, a 580DF40 band-pass filter was used before the transmittedlight is collected by the PMT.

[0216] CE separation. Capillary electrophoresis of the sample wasperformed using a 29-cm long, 20 μm i.d., 150 μm o.d. fused-silicacapillary (Polymicro Technologies, Phoenix, Ariz., USA). Electrophoresisbuffer was a Tris-glycine mixture containing 25 mM Tris and 192 mMglycine at pH 8.3. The injection end of the capillary was set at apositive polarity and the other end installed inside the sheath-flowcuvette was grounded. Sample introduction was performed byelectrokinetic injection at 10 kV for 5 to 10 s unless otherwiseindicated. Separation was performed with an electric field of 330 to 830V/cm.

[0217] Immuno-complex of BPDE-DNA adducts. The incubation conditionswere optimized for short reaction time and stable complex between theBPDE-DNA adduct and its antibody. The incubation was carried out at roomtemperature for 10 min in the dark. The incubation buffer was identicalto the separation buffer except at half the ionic strength. The effectof buffer ionic strength on complex stability was studied by using theTris-glycine buffer at various concentrations.

[0218] Competitive binding of BPDE-DNA adducts. Two oligonucleotides, a16-mer and a 90-mer, were used as probes for competitive immunoassay.They each contained a single BPDE adduct in the middle and both werefluorescently labeled at a 5′ end with a tetramethylrhodamine (TMR).Another adduct standard carrying an identical BPDE-DNA adduct wasprepared. This adduct standard is 16 bases in length and notfluorescently labeled. This BPDE-16 mer competes with the TMR-labeledBPDE-90 mer or TMR-labeled BPDE-16 mer to form complexes with the BPDEantibody. To determine the levels of BPDE-DNA adduct in the A549 cellsexposed to BPDE, the purified genomic DNA from these cells was analyzedand the adducts in the DNA competed with the TMR-labeled BPDE-90 merstandard for binding with the antibody 8E11.

Example 11

[0219] Using CE/LIFP to determine Free solution mobility of DNA adducts.Under free zone electrophoretic conditions, DNA fragments are notseparable when driven by electro-osmotic flow alone because of thesimilar mass-to-charge ratio between DNA fragments. The immunoassaypresented here makes use of an antibody to specifically form a complexwith DNA adducts so that the complex can be separated from the free DNA.The antibody-bound DNA adduct migrates out first and then the unboundDNA. This migration behavior can be expected from equation (8) (Karger,et al., 1989). Eq. (8) predicts the influence of the effective charge(Q), solution viscosity (h) and the radius of an analyte (r) onelectrophoretic mobility (m_(ep)). Both the antibody-bound and unboundDNA adducts are negatively charged. The direction of theirelectrophoretic mobility is opposite to that of the electroosmatic flow(EOF). The decrease in total charge-to-mass ratio after antibody bindingdecreases the mobility of the bound DNA adduct moving back to theinjection end (positive polarity). The net result is a faster migrationdirected towards the detector end (direction of EOF).

m _(ep) =Q/6phr  (8)

[0220] Comparing the two antibodies for their affinity to the BPDE −90mer, antibody 8E11 formed more complex than the antibody 5D11 formed.This may reflect the fact that 8E11 was raised against BPDEmononucleotides and 5D11 was raised against BPDE modified calf thymusDNA. Thus, the 5D11 might be expected to have a higher affinity for longstretches of DNA.

[0221] The longer the capillary column, the more likely that thecomplexes may dissociate during electrophoresis. At a capillary lengthof 60 cm, the amount of detectable antibody-bound DNA adducts wasreduced by approximately 5-fold relative to an identical mixtureseparated on a 30-cm capillary. This reduction may be due to theinstability of the complexes during electrophoretic separation, or maybe caused by adsorption of the complexes on the capillary wall. Theseproblems could be avoided by using a shorter column to carry out theseparation without losing resolution.

[0222] Separation at high field strength also helps to improveresolution. In this Example, at a field strength of approximately 830V/cm (25 kV for 30-cm capillary), the antibody-bound and unbound DNAadducts were baseline resolved in less than 2 minutes, with asignificant improvement in separation efficiency for the antibody-boundDNA adduct. Using Tris-glycine as the separation buffer, Joule heatingwas not excessive at this high field strength as the current generatedwas very low (˜2.2 mA).

[0223] Buffer strength may be varied, with the optimum separation at0.5×Tris-glycine (12.5 mM Tris/96 mM glycine) in this Example. The platecount for the bound and unbound adducts using this buffer condition wascalculated to be 6×10⁵ and 1×10⁶ plates per meter respectively.Incubation time and temperature were also investigated. We observedantibody binding to the DNA adduct at incubation time as short as 1 minand temperature of incubation as low as 0° C. We found that anincubation between 5 and 10 min at ambient temperature was suitable forthe formation of complex and for rapid sample analysis.

[0224] To ensure that the DNA remains in its denatured form, formamidewas added to the incubation buffer to prevent the complementary DNAstrands from being renatured during electrophoresis. Between 2.5 and12.5% (v/v) formamide, the ratio of the bound to unbound adducts wasrelatively constant. Th concentration of formamide should be kept below82.5% (v/v).

Example 12

[0225] Using CE/LIFP in a Competitive assay. Because antibodies arebidentate, each antibody molecule is able to bind with up to two antigenmolecules. One peak in the electropherogram corresponds to the complexbetween one antibody and one DNA adduct. A second peak corresponds tothe complex of one antibody with two DNA adduct molecules. The 1:1 and1:2 complexes between the antibody and DNA adducts are well separated,demonstrating high resolution of the CE system. A competitive assay wasperformed using the TMR-labeled BPDE-16 mer as a probe and the unlabeledBPDE-16 mer as a competitor. As is characteristic of competitive assays,an increase of BPDE-16 mer (unlabeled competitor) corresponds to thedecrease of the complexes between the fluorescent BPDE-16 mer and theantibody.

[0226] The BPDE-90 mer that was fluorescently labeled with TMR was alsoused as a probe to demonstrate competitive immunoassay response with theunlabeled BPDE-16 mer. A similar competitive response was obtained,suggesting that the antibody binds to the BPDE whether it is present inthe 16-mer or the 90-mer oligonucleotides.

Example 13

[0227] Using CE/LIFP to Determination of BPDE-DNA adducts in A549 cells.

[0228] The competitive immunoassay was applied to the determination ofBPDE adducts in A549 cells that were treated with various doses of BPDE.The TMR-labeled BPDE-90 mer was used as the probe and the DNA from A549cells was heat denatured. Increasing amounts of BPDE-DNA adducts wereformed as the cells were incubated with increasing concentrations ofBPDE for 2 hrs. The BPDE-DNA adducts compete with the TMR labeledBPDE-90 mer probe for the antibody binding, resulting in thecorresponding decrease of antibody complexes (peaks 1 and 2) of thefluorescent BPDE-90 mer. As expected one peak corresponding to the 1:1complex between the antibody and the TMR-labeled BPDE-90 mer wasobserved. A second peak attributed to the 1:2 complex of antibody withthe TMR-labeled BPDE-90 mer and the DNA adducts from A549 cells was alsoobserved.

[0229] Using the synthetic BPDE adduct 90-mer as a fluorescent probe andspecific monoclonal antibodies to BPDE-DNA adducts, we demonstrated arapid assay for BPDE-modified DNA in a human lung carcinoma cell line.This approach requires less than 4 min per separation and has excellentresolving power to separate the bound and unbound DNA adducts. The sameapproach may be extended to assays for other types of DNA damage.

[0230] Described in the next series of examples is an assay thatcombines immunological recognition of damaged DNA, capillaryelectrophoresis separation, and laser-induced fluorescence detection(Le, et al., 1998; Xing, et al., 2001). A primary (1°) mouse monoclonalantibody specific for the DNA lesion was used to bind to the DNA lesion.A secondary (2°) anti-mouse IgG antibody that was labeled with afluorescent dye, tetramethylrhodamine (TMR), was used to bind with theprimary antibody. The resulting complex of 2°antibody+1°antibody+damaged DNA was separated using free-zone capillaryelectrophoresis and detected with laser-induced fluorescence. The assaywas used to measure thymine glycol, a typical DNA damage induced byionizing radiation, and to study DNA repair (Le, et al., 1998).Subsequently, the assay was extended to a study of BPDE adducts in DNAfrom human lung carcinoma cells (A549) that were incubated withnanomolar concentrations of BPDE (Xing, et al., 2001).

[0231] Reagents. Unmodified oligonucleotides were synthesized by theDepartment of Biochemistry DNA synthesis laboratory, University ofAlberta, or by Integrated DNA Technologies (Coralville, Iowa). Alloligonucleotides were purified by sequencing polyacrylamide gelelectrophoresis prior to use. Purity of the oligonucleotides wasconfirmed by ³²P-radiolabeling and gel electrophoresis.Tetramethylrhodamine (TMR)-labeled oligonucleotide was synthesized byUniversity Core DNA Services, (University of Calgary, AB).(±)-r-7,t-8-dihydroxy-t-9,10-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene(anti) [(±)-anti-BPDE] was supplied by the National Cancer InstituteChemical Carcinogen Reference Standard Repository (Midwest ResearchInstitute, Kansas City, Mo.). Premixed polyacrylamide/bisacrylamide(19:1) solution was purchased from BioRad Laboratories (Cambridge,Mass.). Enzymes were supplied by Amersham Pharmacia Biotech (Piscataway,N.J.). Monoclonal antibodies 5D11 and 8E11 were purchased from BDPharMingen (San Diego, Calif.). Cell supernatant containing monoclonalantibody E5 (Baan, et al., 1988) was kindly provided by Dr. WilliamWatson, Shell International Chemicals BV, Shell Research and TechnologyCenter, Amsterdam, Netherlands, and was prepared as described by Booth(Booth, et al., 1994). Polyclonal mouse IgG antibody was purchased fromCalbiochem (La Jolla, Calif.). Solvents and other biochemicals weresupplied by Sigma Chemical (St. Louis, Mo.), Fisher Scientific(Pittsburgh, Pa.), or VWR Canlab (Mississauga, ON, Canada).

[0232] Design of probe. In order to imitate DNA damage as it occursnaturally in cellular DNA, we designed a 90-base pair double-strandedoligonucleotide. The desired characteristics of this oligonucleotidewere that it be fluorescently-labeled, contain a known amount of damage,and be long enough to be recognized by a variety of antibodies and otherDNA-binding proteins. The oligonucleotide consists of six overlapping,complementary oligonucleotides of varying lengths that were annealed andligated to form a complete double-stranded 90-mer. The oligonucleotidesequences used in the current study were: oligonucleotide 1:5′-TMR-labeled-CCTTAAGCTTCCTCAACCACTTACCATACTCGAGATT-3′ (SEQ ID NO:2);oligonucleotide 2: 5′-GAGTAT-GGTAAGTGGTTGAGGAAGCTTAAGG-3′ (SEQ ID NO:6);oligonucleotide 5: 5′-GTCATATGCCGCCTCTGA-CCTTCCTAGAATTCCATCC-3′ (SEQ IDNO:8); oligonucleotide 6: 5′-GGATGGAATTCTAGGAAGGTCAG-AGGCGG-3′ (SEQ IDNO:9). The sequence of oligonucleotide 3 and its complementary strand(oligonucleotide 4) may be changed to create a variety of desired damagetypes, typically with a single damaged nucleotide in the middle ofoligonucleotide 3. In the current study the sequences used were:oligonucleotide 3: 5′-CCCATTATGCATAACC-3′ (SEQ ID NO:5); oligonucleotide4: 5′-CATATGACGGTTATGCATAATGGG-AATCTC-3′ (SEQ ID NO:7). The fluorescentLabel (oligonucleotide 1) and damaged nucleotide (oligonucleotide 3) areon the same strand to allow both double- and single-stranded DNAstudies.

[0233] Synthesis of damaged oligonucleotide. (±)-anti-BPDE was used asthe model carcinogen for synthesis of the damaged oligonucleotide. A16-mer with the sequence 5′-CCCATTATGCATAACC-3′ (SEQ ID NO:5) wassynthesized to encourage maximum yield of the BPDE-N² deoxyguanosine(dG) adduct (Margulis, et al., 1993; Funk, et al., 1997). TheBPDE-oligonucleotide reaction was based on the procedure described byMargulis (Margulis, et al., 1993), with slight modifications. The 16-merwas diluted in 20 mM phosphate buffer, (pH 11), containing 1.5%triethylamine, to a concentration of 60 mM in a volume of 400 mL. Afresh 3 mM solution of (±)-anti-BPDE in DMSO was prepared, and 40 mL wasadded to the oligonucleotide solution. This corresponded to aBPDE:oligonucleotide ratio of 5:1. The reaction was carried out at roomtemperature for 20 hours in the dark with gentle shaking.

[0234] Purification of BPDE-oligonucleotide. The components in theBPDE-oligonucleotide reaction mixture were separated usingreversed-phase HPLC. The HPLC system consisted of a Dionex (Sunnyvale,Calif.) AGP1 advanced gradient pump with online-degassing module, eitheran analytical or preparative C18 column, and a Waters (Milford, Mass.)484 tunable absorbance detector in series with a Shimadzu RF-551fluorescence HPLC monitor (Columbia, Md.). The detectors were connectedto a Hewlett Packard Model 35900 multichannel interface (Palo Alto,Calif.), which converted the signals for use by a computer runningChemStation software (Hewlett Packard, Palo Alto, Calif.). Preparativeseparation was carried out on a 10.0×250 mm, 5 mm Luna C18(2)preparative column (Phenomenex, Torrance, Calif.). The reaction productswere initially assessed on the analytical column using a protocoldescribed previously (Margulis, et al., 1993; Cosman, et al., 1990).This procedure employed a linear 0-90% methanol gradient in 20 mM sodiumphosphate buffer (pH 7.0) in 60 min, with a flow rate of 0.75 mL/min. Toreduce separation times for large volumes of the reaction mixture, HPLCpurification of the BPDE-16-mer was carried out in two steps. The firstseparation was under isocratic conditions, using a mobile phase of 70%methanol/30% 20 mM sodium phosphate, pH 7.0 buffer and a flow rate of0.75 mL/min and 3.5 mL/min for the analytical and preparative columns,respectively. Elution of products were monitored in series by theabsorbance detector (wavelength=260 nm for DNA) and the fluorescencedetector (excitation wavelength=343 nm, emission wavelength=400 nm forBPDE). This first separation removed unreacted BPDE as well as thetetrol hydrolysis products. DNA fractions were collected, dried using acentrifugal evaporator, and redissolved in distilled deionized water(ddH₂O). The second separation consisted of a linear 10-40% methanol/20mM sodium phosphate, pH 7.0 buffer gradient in 7.5 min (4%/min) followedby an additional 5 minutes at 40% methanol. This separated theBPDE-oligonucleotide from unreacted oligonucleotide.BPDE-oligonucleotide fractions were collected, dried to remove methanoland redissolved in ddH₂O. The samples were desalted using Sep-Pak C18reversed-phase columns (Waters). The sample was applied to a preparedSep-Pak cartridge, then washed with 10 mL of the following solutions: 25mM ammonium bicarbonate (pH 8.0); 25 mM ammonium bicarbonate/5%acetonitrile; H₂O/5% acetonitrile; H₂O/5% acetonitrile. TheBPDE-oligonucleotide was then eluted with 4×1 mL of H₂O/30%acetonitrile, dried and redissolved in ddH₂O.

[0235] Synthesis and purification of 90-mer oligonucleotides. Prior toligation with the other 5 oligonucleotides, it was necessary tophosphorylate the freshly purified BPDE-16-mer at the 5′-end. Reactionmixtures included: ˜200 pmol of BPDE-16-mer or control 16-mer, 4 mL of100 mM ATP (400 pmol), 1.2 mL of 10× polynucleotide kinase reactionbuffer, and ddH₂O to a total volume of 12 mL. T4 polynucleotide kinase(PNK) was added (1 mL, 6.1 units/mL), then samples were mixed andincubated at 37° C. for 1 hour. After complete reaction, the excess PNKwas heat denatured at 70° C. for 10 minutes. The 16-mers were then mixedwith the TMR-labeled 37-mer and the other 4 oligonucleotides so that allwould be in 2:1 excess over the 16-mers. 5×DNA ligase buffer was addedto a final concentration of 1× and the mixture was heated in a waterbath to 70° C. for 10 minutes, then allowed to cool over several hoursto room temperature. DNA ligase was added (2 mL, 8.5 Weiss units/mL) andthe sample incubated overnight at 16° C.

[0236] Purification of the BPDE and control ligation products wasachieved using preparative, 7.5% native polyacrylamide gelelectrophoresis (PAGE). Blectrophoresis was carried out at 600 V for 6hours with a water cooling core to prevent denaturation of the ligationproducts. The bands were visualized by brief exposure to ultravioletlight, causing the TMR label to fluoresce, and cut from the gel. The gelslices were crushed and soaked to elute the products overnight in 0.3 Msodium acetate, pH 5.2 on a rotary shaker protected from light. Afterelution, polyacrylamide fragments were removed from solution usingfilter units prepared in the lab. The solution was passed throughsilanized glass wool followed by GF/C glass microfibre filter paper(Whatman). The samples were then extracted and back-extracted with equalvolumes of phenol/chloroform/isoamyl alcohol (25:24:1) followed bychloroform/isoamyl alcohol (24:1). Oligonucleotides were precipitated byadding MgCl₂ to 10 mM and 3 volumes of ice-cold 95% ethanol and, thenplaced at −20° C. overnight. The following day samples were centrifugedfor 45 minutes at 14000 rpm and 4° C., supernatant was removed, and thepellets were washed once with 95% ethanol. Samples were againcentrifuged for 10 minutes, dried and redissolved in ddH₂O. UV-Visabsorbance scans were performed on the resulting oligonucleotidesolutions to determine concentration as well as to confirm the presenceof the TMR dye and BPDE moiety.

[0237] Instrumentation for analysis of ligation products, was the sameas described in examples 11-13. with the following modification. Thesystem was equipped with an auxiliary microscope to assist in thealignment of the optics. The microscope was used to visualize theposition of the laser beam with respect to both the sample flow throughthe capillary and the collection optics, represented by a light-emittingdiode (LED) positioned behind the pinhole in the collection assembly.Alignment was achieved by initially fixing the position of thecollection assembly, then adjusting the capillary and laser-focusingobjective using X-Y-Z translation stages. The angle of thefluorescence-collecting objective and the position of the collectionassembly were also adjustable for optimization of alignment.

[0238] Samples were electrokinetically injected into the capillary byapplying an injection voltage of 10000 V for 5 seconds. The separationwas carried out at room temperature with a separation voltage of 20000V. The running buffer used was 1×Tris-glycine (25 mM Tris, 250 mMglycine), pH 8.3. The capillary was washed approximately every 5-10injections with 0.1 M NaOH (applied by syringe for 1 min) followed byelectrophoresis using 1× Tris-glycine, pH 8.3 for 7 minutes. The initialvoltage was kept low to prevent excessive joule heating in thecapillary. As the running buffer replaced the NaOH in the capillary,current decreased allowing the running voltage to be gradually increasedto 20000 V for the final 5 minutes of the reconditioning period. Allcapillary electrophoresis data were analyzed using Igor Pro software(version 3.1, WaveMetrics Inc., Lake Oswego, Oreg.).

[0239] Characterization of BPDE and control 90-mers. Prior to analysis,90-mer samples were diluted to appropriate concentrations in runningbuffer (1× Tris-glycine, pH 8.3). The 90-mer products were analyzedeither in their native form or their denatured, single-stranded form.Denaturation of the 90-mers was achieved by heating the samples at 100°C. for 10 minutes in a heating block, then transferring directly to iceto prevent reannealing. After cooling, the samples were brieflycentrifuged in a microcentrifuge to collect condensation from the sideof the tube, then gently mixed to ensure a homogenous solution. Totalsample volume was typically 20 mL, which allowed for convenientinjection into the capillary. For experiments involving antibodies,fresh dilutions of antibody stock solutions were prepared immediatelybefore analysis and kept on ice. After addition of antibody to the90-mer solution, the sample was gently vortexed to ensure completemixing.

[0240] Treatment of A549 cells with BPDE. A human lung carcinoma cellline (A549) was incubated with BPDE to produce DNA adducts in genomicDNA. The cell line was maintained in DMEM/F12 medium (Gibco BRL,Gaithersburg, Md.) supplemented with 10% fetal bovine serum. The cellswere seeded at 1×10⁵ cells per plate and maintained at 95% humidity and5% CO₂ for 20 hours prior to the addition of BPDE. Old culture mediawere removed from each culture plate and the cells were washed twicewith phosphate buffered saline (PBS). Media containing BPDE at variousconcentrations (0, 2.5, 5, and 10 mM final concentration) were added tothe designated plates. The cells were further incubated in the mediacontaining BPDE for 2 hours. The cells were then washed with PBS priorto the addition of DNAzol lysis reagent (Gibco BRL) to facilitate celllysis and DNA extraction. Subsequent steps involved a 99.9% ice coldethanol precipitation and a 70% cold ethanol wash to purify the genomicDNA. The final DNA pellet was dissolved in distilled deionized water(ddH₂O) and DNA concentration was measured at OD₂₆₀ using ddH₂O as ablank.

[0241] Competitive assay for BPDE-DNA adducts. The DNA samples from theA549 cells were analyzed for BPDE-DNA adducts by competitive assay usingthe TMR-labeled 16-mer or 90-mer oligonucleotides as probes. Mixturescontaining 60 nM of the oligonucleotide probe, 0.4 mg/mL of mousemonoclonal antibody 8E11, and 80 mg/mL of the DNA from A549 cells wereincubated in 20 mL of tris-glycine buffer (25 mM tris and 200 mMglycine, pH 8.3) at room temperature for 30 min. These were subjected toCE/LIF analysis to detect both antibody-bound and unbound fluorescentprobes.

Example 14

[0242] Using CE/LIF to determine Affinity interactions of BPDE-90-merswith a monoclonal antibody. The purification of BPDE-16-meroligonucleotide, the synthesis and purification of BPDE 90-mer ligationproducts, and the characterization of the BPDE 90-mer ligation productswere carried out as known to those skilled in the art.

[0243] Preliminary experiments using monoclonal antibody 8E11demonstrated that the specific antibody bound to the BPDE-90 mer, notthe control 90 mer. These results were obtained by first denaturing the90-mers (5×10⁻⁹ M), then adding 8E11 antibody to a final concentrationof 20 mg/mL and incubating for 10 min at room temperature (21° C.). Thesame fluorescence intensity scale was used for both 90-mers for ease ofcomparison. For the mixture of the BPDE 90-mer and 8E11, an additionalpeak was present in the electropherogram with a migration time ofapproximately 3.0 min. This peak represented the complex between theantibody and single-stranded BPDE-DNA, and was well-resolved from thedenatured 90-mer peak at 4.1 min. When comparing the fluorescent signalsbetween runs, the total area of the two peaks for the mixture of BPDE90-mer and 8E11 was very similar to the area of the peak for the BPDE90-mer alone. The formation of an antibody-DNA complex was not observedwith the control 90-mer, indicating a specific interaction of theantibody with the BPDE 90-mer.

[0244] The effect of incubation time and temperature on complexformation were investigated using the same concentrations of BPDE 90-mer(5×10⁻⁹ M) and 8E11(20 mg/mL). For incubations carried out at both roomtemperature and on ice, the interaction did not change significantlybetween 1 min and 20 min. At room temperature, the complex was stableafter 45 min. For incubation on ice, the complex decreased slightlyafter 45 min when compared to the 20 min incubation. In general, roomtemperature incubations with 8E11 resulted in more stable andreproducible complex formation than incubations on ice. This result isexpected since the recommended temperature for conventional immunoassaysusing 8E11 is 37° C. (Santella, et al., 1984; Hsu, et al., 1995), andmost inumunochemical procedures require incubation temperatures ofeither 37° C. or room temperature. Based on these results, furtherexperiments with 8E11 antibody were carried out at room temperature. Anincubation time of 5 min was chosen for ease of sample preparation andanalysis.

[0245] Overnight incubations resulted in a decrease of the DNA-antibodycomplex, as well as a reversion to the doublet shape for the free 90-merpeak. This result suggests that 90-mer samples left overnight tended tore-anneal to the double-stranded form, causing dissociation of theDNA-antibody complex. This also implies that the affinity of 8E11 fordouble-stranded BPDE-DNA is less than for single-stranded BPDE-DNA.

[0246] The difference in affinity of 8E11 antibody between single- anddouble-stranded BPDE-modified DNA was further confirmed by comparing itsbinding with heat-denatured BPDE 90-mer and native BPDE-90 mer (20 mg/mL8E11). Antibody-oligonucleotide complex formation was approximately 6fold higher for the denatured single-stranded 90-mer than the nativeform. Thus, denaturation of samples by heat before incubation withantibodies was retained for further experiments.

Example 15

[0247] Using CE/LIF to Determination of specific antibody using BPDE-90mer as a probe. An application of the fluorescent BPDE-90 mer probe wasdemonstrated for the determination of anti-BPDE antibody. Calibrationfrom the analyses of mixtures containing different amounts of 8E11 and aconstant concentration of the detatured BPDE-90 mer probe (5×10⁻⁹ M)were made. A DNA-antibody complex peak was observed with 8E11concentrations as low as 0.1 mg/mL. This concentration corresponds to0.7×10⁻⁹ M (or 0.7 nM) assuming a molecular weight of approximately150,000 for the antibody 8E11. The concentration of BPDE-90 mer (5 nM)was in excess and the formation of its complex with the antibody was notcomplete. The amount of the complex increased at higher concentrationsof 8E11, up to 10 mg/ml (7 nM). At this concentration complex formationappeared to reach saturation, since further increase of antibodyconcentrations did not increase the proportion of 90-mer bound to 8E11.

Example 16

[0248] Using CE/LIF for Screening for anti-BPDE antibodies using thefluorescent BPDE-90mer probe. The fluorescent BPDE-90 mer probe wasfurther used to screen for specific binding proteins, with 3 antibodiesas model protein analytes. Monoclonal antibodies 8E11, 5D11 and E5 areall specific for BPDE-modified DNA. A comparison between theseantibodies was conducted to determine differences in their reactivity tothe BPDE 90-mer standard as well as their behavior in the capillaryelectrophoresis system. Conditions used for sample preparation wereidentical to earlier experiments: heat denaturation of the 90-mer at100° C. for 10 min, cooling on ice, then incubation with antibody atroom temperature for 5 min before injection. Polyclonal mouse IgG wasused as a negative control since it is essentially the same molecularstructure (isotype) as the monoclonal antibodies but is not expected toreact with the BPDE 90-mer. The BPDE 90-mer probe concentration wasfixed at 5×10⁻⁹ M and the antibodies were added in varying amounts. Allthree monoclonal antibodies reacted with the 90-mer probe, with 8E11giving the highest formation of complex. The negative control showed avery slight reactivity but was insignificant compared to the otherantibodies, even at concentrations up to 40 mg/mL.

[0249] Antibodies 8E11 and E5 were found to bind specifically to theBPDE adduct. No cross-reactivity with the unmodified control 90-mer wasobserved for either 8E11 or E5. The antibody 5D11 showed slightcross-reaction with undamaged DNA. When incubated with 20 μg/mL 5D11,the control 90-mer formed a peak corresponding to antibody complex, withabout 2.1% of total peak areas as compared with the BPDE 90-mer. Thisnon-specific interaction between 5D11 and undamaged DNA is in agreementwith previous studies (Santella, et al., 1984) that have demonstratedcross-reactivity, and is a result of its being raised against afull-length BPDE-DNA antigen. Both 8E11 and E5 were raised againstBPDE-guanosine monomers conjugated to carrier proteins (Baan, et al.,1988; Santella, et al., 1984) and therefore do not recognize undamagedDNA.

[0250] The incomplete binding of the DNA damage probe with theantibodies (up to 50% of binding) is probably because the probe is amixture of several BPDE-90 mer isomers. The stereochemistry of theBPDE-N²-dG adduct could be important to its binding with specificantibodies. In the preparation of the BPDE-modified 16-mer,(±)-anti-BPDE was reacted with the oligonucleotide. The covalent bondthat forms between BPDE and guanosine may be either cis- ortrans-relative to the hydroxyl group on the adjacent carbon atom.Therefore, there may be as many as four different configurations of theBPDE 16-mer: (+)-trans, (+)-cis, (−)-trans, and (−)-cis (2). Thereaction protocol was designed to minimize the formation of cis-adducts(Funk, et al., 1997), but a mixture of (+)-trans and (−)-trans adductswith a small amount of cis adducts would be expected in the BPDE 16-merreaction products (Cosman, et al., 1990). Because these stereoisomerswere pooled together after purification by HPLC and before the ligationreaction, the 90-mer product would also contain these configurations.The advantage of this mixture is that it more accurately represents thespectrum of damage that would occur in human DNA samples. Thedisadvantage is that BPDE-DNA antibodies exhibit different affinitiesfor these stereoisomers (Hsu, et al., 1995). In competitive inhibitionstudies using BPDE-modified 11-mers, Hsu (Hsu, et al., 1995)demonstrated a lower affinity for the (−)-trans-anti-BPDE-N²-dG adductthan for the (+)-trans-anti-BPDE-N²-dG adduct. For antibodies 8E11 and5D11 this lower affinity was 66% and 20% of the (+)-trans adduct,respectively. Both antibodies exhibited much lower affinities for thecis adducts compared to the (−)-trans adduct. Since the 90-mer containeda combination of both trans adducts, the stereospecific difference inaffinity may in part be responsible for the differences in complexformation observed for these antibodies. The presence of differentBPDE-90mer isomers may also contribute to the observed incompletebinding. The other possible reason for the incomplete binding is thepresence of residual oligonucleotides that do not contain BPDE andtherefore, do not bind to the antibodies.

[0251] In addition to the isomer-specific reactivities, Hsu (Hsu, etal., 1995) showed a difference in affinity between 8E11 and 5D11 whenconsidering only the (+)-trans adduct. 8E11 was approximately 7 timesmore sensitive than 5D11 for the very short 11-mer oligonucleotide. Forfull-length heat-denatured BPDE-DNA, the two antibodies were almostidentical. This difference is likely due to the antigens against whichthese antibodies were raised: BPDE-N²-dG mononucleotide for 8E11,full-length BPDE-DNA for 5D 11. 5D 11 may require a longer sequence ofDNA surrounding the damaged site for binding which would not be presentin the 11-mer. Given these results one might predict that for DNA ofintermediate length (90 bases), 8E11 would still have a higher affinitythan 5D 11, but to a lesser extent. These results are consistent withprevious findings, which indicates that monoclonal antibody 8E11 islikely the best choice for detecting BPDE-damaged DNA using thecapillary electrophoresis/laser-induced fluorescence assay.

Example 17

[0252] Using CE/LIF and the BPDE-DNA probe in a competitive assay forBPDE-DNA adducts in cells. The 90-mer probe described herein has manypotential uses in DNA damage research. It enables the investigation ofalternative assay methods, including CE-based competitive immunoassays(Tao, et al., 1996; Ye, et al., 1998; Lam, et al., 1999; Wan, et al.,1999) using the probe as a fluorescent probe (competitor). This approachis based on competition between damaged DNA and the fluorescent probefor the binding sites of a limited amount of antibody. With little or nodamaged DNA in a sample, the probe achieves maximum complex formationwith the antibody. As the amount of damaged DNA in the sample mixtureincreases, the probe is displaced from the antibody. This would resultin an increase in the free probe peak and a decrease in theprobe-antibody complex peak. This method has been demonstrated by usingoligonucleotide and genomic DNA containing BPDE-damaged sites (Tan, etal., 2001). Electropherograms from the analysis of BPDE-DNA adducts inA549 cells that were incubated with 2.5, 5, and 10 mM BPDE for 2 hr werecollected. Again, increasing amounts of BPDE-DNA adducts were formed asthe cells were incubated with increasing concentrations of BPDE. TheBPDE-DNA adducts compete with the TMR labeled BPDE-DNA adduct probe forthe antibody binding, resulting in the corresponding increase of theunbound probe (peak 3) and decrease of antibody complexes (peaks 1 and2) of the fluorescent probe. This analysis requires less than 4 min perseparation and has excellent resolving power to separate the bound andunbound DNA adducts. The same approach may be extended to assays forother types of DNA damage.

[0253] Another important aspect of the probe's design is the flexibilityto substitute different damage types in the molecule with relative ease.The sequences of the two center oligonucleotides may be changeddepending on the desired modification. By inserting these differentdamaged oligos, a variety of DNA damage detection systems can beinvestigated using the corresponding damage probe and CE/LIF. Thetechnique itself combines specific recognition with high sensitivitydetection, minimal sample preparation, and fast analysis times (5minutes per run).

Example 18

[0254] Using CE/LIF to Determine Stoichiometry of antibody binding withTMR-BPDE-16-mer (16mer*) oligonucleotide. In this example CE/LIF is usedto determine the binding stoichiometry of DNA adducts with antibodies.Advantage is taken of the fact that both size and charge of themolecules contribute to CE separation. If additional charges can beintroduced to the complex due to binding, then the separation of themultiple complexes becomes possible. Fluorescent oligonucleotide probesthat contain a single adduct which can be recognized by an antibody weredesigned. These probes introduce large mobility changes to the antibodywhen bound to the probe because of the highly negative charge of theprobe. With these probes, we are able to study the binding stoichiometrybetween oligonucleotides and the antibody. DNA adducts of benzo[a]pyrenediol epoxide (BPDE) were looked at in this example. Available monoclonalIgG antibody has a high affinity for BPDE-DNA adducts, allowing detailedinformation on binding stoichiometry between the antibody and the DNAadducts to be obtained. This example provides direct information onantibody binding stoichiometry.

[0255] Reagents: Oligonuleotides were synthesized by the Department ofBiochemistry DNA synthesis laboratory, University of Alberta, or byIntegrated DNA Technologies (Coralville, Iowa). All oligonucleotideswere purified by sequencing polyacrylamide gel electrophoresis prior touse. Purity of the modified oligonucleotieds was confirmed by gelelectrophoresis and ³²P-postlabeling. Tetramethylrhodamine (TMR)-labeledoligonucleotide was synthesized by University Core DNA Services,(University of Calgary, AB).(±)-r-7,t-8-dihydroxy-t-9,10-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene[(±)-anti-BPDE] was supplied by the National Cancer Institute ChemicalCarcinogen Reference Standard Repository (Midwest Research Institute,Kansas City, Mo.). Mouse monoclonal antibody 8E11 was purchased from BDPharMingen (San Diego, Calif.). Polyclonal rabbit IgG antibody waspurchased from Calbiochem (La Jolla, Calif.). Solvents and otherbiochemicals were supplied by Sigma (St. Louis, Mo.), Fisher Scientific(Pittsburgh, Pa.), or VWR Canlab (Mississauga, Ontario).

[0256] Synthesis of BPDE-DNA adducts: Two 16-mers with the sequence5′-CCCATTATGCATAACC-3′ (SEQ ID NO:5) were synthesized and reacted withBPDE to yield the BPDE-N² deoxyguanosine (dG) adduct (Margulis, et al.,1993; Funk, et al., 1997). One of the 16-mer oligonucleotides waslabeled with TMR at the 5′ end, and the other was not labeled. Theformation of BPDE-oligonucleotide was based on the procedure describedby Margulis (Margulis, et al., 1993) with slight modifications. The16-mer was diluted in 20 mM phosphate buffer (pH 11) containing 1.5%triethylamine, to a concentration of 60 mM in a volume of 400 mL. To theoligonucleotide solution was added 40 mL 3 mM BPDE in DMSO. Thiscorresponded to a BPDE:oligonucleotide ratio of 5:1. The reaction wascarried out at room temperature for 20 hours, in the dark with gentleshaking. The double stranded TMR-BPDE-90-mer and BPDE-90-mer wereconstructed through ligation of BPDE-16-mer with five otheroligonucleotides.

[0257] Purification of TMR-BPDE-oligonucleotide: The components in theTMR-BPDE-oligonucleotide reaction mixture were separated usingreversed-phase HPLC. The HPLC system consisted of a Dionex (Sunnyvale,Calif.) AGP1 advanced gradient pump with online degassing module, ananalytical C18 column, a Waters (Milford, Mass.) 484 tunable absorbancedetector in series with a Shimadzu (Tokyo, Japan) RF-551 fluorescencedetector. The detectors were connected to a Hewlett Packard (Palo Alto,Calif.) Model 35900 multichannel interface, which converted the signalsfor use by a computer running ChemStation software (Hewlett Packard).The analyses were performed using a Luna C18(2) analytical column(4.6×250 mm, 5 mm, Phenomenex, Torrance, Calif.). The reaction productswere purified initially using a gradient elution. This procedureemployed 10 mM sodium phosphate buffer (pH 7.0) and acetonitrile. Theacetonitrile content was initially 10%, linearly ramped to 15% duringthe first 20 min, and then kept at 15% for 5 min. After theBIPDE-modified oligonucleotides were eluted and collected, the columnwas washed using 50% acetonitrile for 20 min to remove unreacted BPDEand its metabolites. The flow rate was 1.0 mL/min. Elution of productswere monitored in series by an absorbance detector (wavelength=260 nmfor DNA) and a fluorescence detector (excitation wavelength=535 nm,emission wavelength=580 nm for TMR). The fractions containing theBPDE-oligonucleotide were pooled and further purified to remove anyunmodified oligonucleotides. This second HPLC purification step wascarried out using an isocratic elution with 10 mM phosphate buffer (pH7.0), containing 12.5% acetonitrile as mobile phase. Other HPLCconditions were the same as described above. The collected fractionswere concentrated by pressurized air under room temperature anddissolved in distilled deionized water (ddH₂O). The purifiedBPDE-oligonucleotides were evaluated using HPLC under isocratic elutionconditions, and showed good purity of above 95%. The concentration ofthe final TMR-BPDE-16-mer was estimated using absorbance at 260 nm.

[0258] Instrumentation for analysis of the DNA-BPDE adducts: Analysisand characterization of the DNA-BPDE adducts was carried out using alaboratory-built capillary electrophoresis laser induced fluorescence(CE/LIF) system as described in examples 13 on.

[0259] Samples were electrokinetically injected into the capillary byapplying an injection voltage of 15 kV for 5-10 seconds. The separationwas carried out at room temperature with a separation voltage of 15 kV.The running buffer were either 1× tris-glycine (25 mM Tris, 192 mMglycine, pH 8.3) or 0.5×tris-glycine (12.5 mM tris and 96 mM glycine, pH8.3). The capillary was washed approximately every 3 injections with0.02 M NaOH electrophoretically at 15 kV for 7 min followed byelectrophoresis using water and the running buffer for 7 min each. Allcapillary electrophoresis data were analyzed using Igor Pro software(version 3.1, WaveMetrics Inc., Lake Oswego, Oreg.).

[0260] Complex formation of TMR-labeled BPDE-DNA adducts and antibody:TMR-BPDE-16-mer and TMR-BPDE-90-mer samples were diluted to appropriateconcentrations in running buffer (tris-glycine, pH 8.3). The doublestranded BPDE-90-mer was denatured by heating at 95° C. for 5 min in aheating block. It was then placed on ice to prevent reannealing. Aftercooling, the samples were briefly centrifuged in a microcentrifuge(Z233M, Hermle) to collect condensation from the side of the tube, thengently mixed to ensure a homogenous solution. TMR-BPDE-16-mer wassynthesized as a single-stranded oligonucleotide. Appropriate dilutionsof antibody stock solutions were prepared immediately before use andkept on ice. After addition of antibody to the BPDE-oligonucleotidesolutions, the samples were gently vortexed to ensure complete mixingand incubated at the room temperature for 5-10 min, then analyzed byCE/LIF. The total sample volume was typically 20 mL.

[0261] Simultaneous binding of two BPDE-DNA adducts to the antibody:Both TMR-labeled and unlabeled BPDE-adducts were allowed to bind withthe antibody. The freshly diluted BPDE-16-mer (16mer) andTMR-BPDE-16-mer (16mer*) solutions were mixed together before additionof the antibody. Concentrations of the TMR-BPDE-16-mer and the antibodywere kept constant at 9.6 nM and 33.3 nM, respectively, and theunlabeled BPDE-16-mer was varied from 2.5 nM to 5.0 mM. The samplemixtures were incubated for 5 min at room temperature, then analyzed byCE/LIF. Similarly, the antibody was added to mixtures of theTMR-BPDE-16mer and BPDE-90mer to study the competitive binding andligand exchange.

[0262] A series of electropherograms from CE/LIF analyses of mixturescontaining 24 nM TMR-BPDE-16-mer (16mer*) and varying concentrations ofmouse monoclonal antibody (Ab) to BPDE, from 0.5 mg/mL to 16.0 mg/mLwere collected. In these mixtures, 2 complexes between the Ab and theTMR-labeled BPDE-16-mer (16mer*) can be expected as follows:

Ab+16mer*=Ab(16mer*)+Ab(16mer*)₂  (9)

[0263] The free (unbound) 16mer* gives a peak that was well resolvedfrom the two complexes. The unbound 16mer* oligonucleotide and theantibody-bound 16mer* complexes are negatively charged. Under the freezone CE separation conditions, the direction of their electrophoreticmobility (μ_(ep)) is opposite to that of the electroosmotic flow (EOF).Among the three fluorescent species the unbound 16mer* has the highestnegative effective charge. It has the highest electrophoretic mobilitytowards the positive (injection) end, and thus the longest migrationtime (3.4 min). When the 16mer* binds to an antibody molecule, thereduction of the effective charge results in a smaller electrophoreticmobility and thus a shorter migration time (2.1 min). When the Ab bindsto two 16mer* molecules, the charge density of the complex is betweenthose of the Ab(16mer*) complex and the unbound 16mer*. Therefore, asmaller mobility shift is expected. Indeed, we observed the Ab(16mer*)₂complex at 2.4 min. The two complexes are well resolved from each other,(resolution of 1.55).

[0264] It is evident that formation of the two complexes depends on therelative concentrations of the TMR-BPDE-16-mer oligonucleotide (16mer*)and the antibody. The intensity of Ab(16mer*) complex (1:1stoichiometry, peak 1) increases with increasing concentration of theantibody, whereas the Ab(16mer*)₂ complex (1:2 stoichiometry, peak 2)reaches a maximum at an antibody concentration of 1.0 μg/mL and thendecreases gradually with increasing concentrations of the antibody.

[0265] The primary complex of the 1:1 stoichiometry increases withincreasing concentration of antibody. When the concentration of theantibody is 8-16 ,μg/mL, approximately 80-85% of the total 16mer*oligonucleotide is present as the primary complex with the antibody (1:1stoichiometry). Formation of the secondary complex, Ab(16mer*)₂ (1:2stoichiometry) is favored at lower concentrations of the antibody, whenthe 16mer* oligonucleotide is in excess. The amount of the secondarycomplex reaches a maximum at the antibody concentration of 1 μg/mL, when50% of the total 16mer* is present as the secondary complex. As themolecular weight of the mouse monoclonal antibody IgG is approximately150,000 Da, an antibody concentration of 1 μg/mL is approximately 7 nM.Although the Ab(16mer*)₂ complex is observed throughout the entireantibody concentration range studied, from 0.02 μg/mL (˜0.14 nM) to 16μg/mL (˜100 nM), it is predominant only when the antibody concentrationis below 2 μg/mL (˜14 nM) and is lower than the concentration of the16mer* (24 nM). At similar concentrations of antibody (4 μg/mL or 27 nM)and the 16mer* (24 nM), the primary complex dominants, suggesting thatthe complex with one binding site is preferred.

[0266] To further study the secondary complex, varying concentrations ofthe 16mer* (2.4-19.2 nM) were mixed with a fixed concentration of theantibody (0.4 μg/mL or ˜2.7 nM), and the mixtures were analyzed usingCE/LIF. Fluorescence intensity of the secondary and primary complexes asa function of the concentration of the fluorescently labeled BPDE-16-mer(16mer*) were measured. At a lower concentration of the 16mer* (2.4 nM)than the antibody concentration (2.7 nM), the peak intensity of theprimary complex is comparable to that of the secondary complex. When the16mer* concentration is higher (4.8-19.2 nM) than the antibodyconcentration (˜2.7 nM), the secondary complex dominates. Thefluorescence intensity of the secondary complex increases withincreasing amounts of the 16mer* until it reaches a plateau atapproximately 17 nM when the amount of the antibody presumably becomesthe limiting factor.

[0267] Most previous studies on immunoassays were not able to addressthe binding stoichiometry although multiple complexes might have formed.The present study clearly shows the formation of primary and secondarycomplexes. The behavior may be qualitatively explained by a two bindingsite model. The mouse monoclonal antibody is an IgG, which has twospecific binding sites for antigen (hapten), in this case, BPDE-16-meroligonucleotide. When an antibody is in excess of antigen, there aremore available binding sites for antigen, thus most complex exists asthe primary complex (1:1 stoichiometry). Only when the binding sites arelimited as in the case of lower antibody concentration, the secondarycomplex (1:2 stoichiometry) dominates. These results suggest thatbinding to the second sites of antibody is less favored probably becauseof possible steric hindrance by the binding on the first site.

Example 19

[0268] Using CE/LIF to Determine the Binding of the antibody withTMR-BPDE-90-mer (90mer*): To confirm the preferential binding to thefirst site and the possible hindrance to the secondary binding, wefurther compared binding of the antibody with a TMR-BPDE-90-mer (90mer*)oligonucleotide. The 90mer* is approximately 28,000 Da, about five timeshigher than the 16mer* (˜5,000 Da). Eelectropherograms from CE/LIFanalyses of samples containing 50 nM 90mer* incubated with varyingconcentrations of mouse monoclonal antibody, 0-20 μg/mL (0-140 nM) werecollected. In the absence of antibody, the 90mer* gave a single peak atmigration time of 3.2 min. After incubation with the antibody, twoadditional peaks are present in the electropherogram with migrationtimes of 2.41 and 2.60 min. These peaks represent the complexes betweenthe antibody and single stranded 90mer* oligonucleotide. Because of theelectrophoretic mobility shift, they are well resolved from the unbound90mer*. Despite the small difference in migration time between the twocomplexes (0.19 min), these two complexes are baseline resolved. Thefirst peak is due to the primary complex of the antibody and DNA adductwith 1:1 stoichiometry, and peak 2 is due to the secondary complex of1:2 stoichiometry.

[0269] The binding of the anti-BPDE antibody with TMR-BPDE-90-merappears to be weaker than the binding with TMR-BPDE-16-mer.Approximately 60% of the total 90mer* (50 nM) formed complex with theantibody when the concentration of antibody was ˜140 nM. This percentageis lower than that for the 16mer* where 80-85% of the total 16mer* (24nM) complexed with the antibody (50-100 nM). This is not surprisingconsidering that the antibody (mouse monoclonal, 8E11) was raisedagainst BPDE-adduct of mononucleotide and perhaps has a higher affinityto shorter stretches of DNA.

[0270] The ratio of the primary and secondary complexes was also foundto depend on the relative concentrations of the antibody and the 90mer*,similar to that shown in the 16mer* experiments described above.However, the secondary complex of the antibody with the 90mer* is lessstable than the secondary complex of the same antibody with the 16mer*.The primary complexes of the antibody with both the 90mer* and the16mer* are much more stable. The results support our suggestion that theprimary binding is stronger than the secondary binding. Despite theidentical sequence and structure of the two binding sites of themonoclonal IgG antibody, it is likely that the secondary binding isaffected by the primary binding on the first site. This may be caused bya conformational change of the antibody structure after binding to onesite. Alternatively, steric hindrance of the molecule bound to one siteof the antibody may affect the binding of another molecule on the secondsite of the same antibody molecule. A comparison of our results on the16mer* and 90mer* indicates that the secondary binding with the larger90mer* is less favored. These results suggest that steric hindranceplays an important role in the formation of the secondary complexbetween an antibody and an antigen.

Example 20

[0271] Using CE/LIF to measure competitive binding of antibody withTMR-BPDE-16-mer and unlabeled BPDE-16-mer: The commonly used competitiveimmunoassays are based on competitive binding of two ligands to alimiting amount of antibody. Typically, one ligand is labeled and isused as a detection probe. The target analyte is usually the unlabeledligand. In the traditional competitive assay format, the analyte(unlabeled ligand) can only be indirectly determined through monitoringthe relative intensity of signals produced from the labeled ligand. Inthis study, It was found that the primary and secondary complexes of theantibody with the ligands can be separated. Thus, we decided to furtherstudy the binding of multiple ligands to the antibody, with apossibility of developing new approaches to binding assays.

[0272] Electropherograms from CE/LIF analyses of mixtures containing 33nM antibody, 35 nM TMR-BPDE-16-mer (16mer*), and varying concentrations(0, 0.05, 0.5 and 5.0 mM) of unlabeled BPDE-16-mer (16mer) werecollected. In the absence of the unlabeled BPDE-16-mer, the primarycomplex [Ab(16mer*)] dominates and its fluorescence intensity is muchhigher than that of the secondary complex [Ab(16mer*)₂].

[0273] While the primary complex decreases with increasingconcentrations of the competing unlabeled 16mer, a typical competitiveimmunoassay behavior, the secondary complex increases initially withincreasing concentrations of the 16mer. The latter behavior is uniqueand has not been reported previously with competitive immunoassays. Thisobservation can be explained as following.

[0274] The present CE/LIF technique allows the separation of twoantibody complexes when a labeled ligand (L*) (e.g., 16mer*) is mixedwith the antibody.

Ab+L*=AbL*+AbL* ₂  (10)

[0275] To allow for detection of an unlabeled ligand (L), competitiveimmunoassay approaches rely on the competition of unlabeled ligand (L)with the labeled ligand (L*) for the limiting amount of antibody (Ab).Following species may be formed:

AbL*+L=AbL+L*  (11)

AbL*+L=AbL*L  (12)

AbL* ₂ +L=AbL*L+L*  (13)

AbL* ₂+2L=AbL ₂+2L*  (14)

[0276] The species that are fluorescent and can be detected include thefree ligand L*, the primary complex AbL*, and the secondary complexesAbL*L and AbL₂*. It is clear that the end products are a redistributionof L* and L in the complexes although L* and L compete for the sameantibody.

[0277] The relative changes of the primary and secondary complexes withvarying concentrations of unlabeled 16mer when it is incubated with 9.6nM TMR-labeled 16mer* and 33 nM (5 μg/mL) antibody were measured. Whenthe concentration of the unlabeled 16mer is below 10 nM, the primarycomplex [Ab(16mer*)] dominates and its amount remains almost constantwith increasing concentration of the unlabeled 16mer up to 10 nM. Thetotal ligand concentration (16mer* and 16mer) is lower compared withthat of the antibody, and there are excess antibody binding sitesavailable. Thus, no significant competition between the two ligandstakes place. Increasing the competing 16mer concentration from 10 nM to200 nM result in the reduction of the primary complex in a mannersimilar to that commonly observed for conventional competitive assay.Further increase of the unlabeled 16mer concentration to above 250 nMresults in disappearance of the primary Ab(16mer*) complex, probably dueto the displacement of the 16mer* by the excess 16mer:

Ab(16mer*)+16mer=Ab(16mer)+16mer*  (15)

[0278] The secondary complexes initially increases with increasingconcentration of the unlabeled 16mer, and reaches a maximum when theconcentration of the 16mer is 100 nM. This behavior is different fromthat of traditional competitive immunoassays where only the mixture ofantibody complexes are commonly detected and the separation of the 1:1and 1:2 complexes is not available for examination. Further increase ofthe 16mer concentration (100-1000 nM) results in a gradual decrease ofthe secondary complex. Only at this high concentration range (100-10000nM) of the competing 16mer was the competitive immunoassay behaviorobserved.

[0279] When the secondary complex and the unlabeled BPDE-16-mer areplotted using logarithm scales, two linear lines are observed. A linearcorrelation coefficient of 0.97 and a positive slope are observedbetween the secondary complex and the concentration of the 16mer below100 nM. This region probably corresponds to redistribution of theligands. Above 100 nM, a linear correlation of 0.997 and a negativeslope are observed. This may indicate competition with limited amount ofantibody. These two linear lines clearly distinguish the competition andredistribution, and can be used as quantitative curves for differentconcentration zones.

[0280] It is expected that two types of the secondary complexes could beformed: one antibody bound to two TMR-BPDE-16-mer [Ab(16mer*)₂] and oneantibody bound to one TMR-BPDE-16-mer and one BPDE-16-mer[Ab(16mer*)(16mer)]. These two secondary complexes of 16-mer cannot beseparated since there is only slight difference between the TMR-labeledand unlabeled 16-mer oligonucleotides. However, the two secondarycomplexes can be observed using a different competing oligonucleotide asdescribed below.

Example 21

[0281] Using CE/LIF in competitive binding studies betweenTMR-BPDE-16-mer and unlabeled BPDE-90-mer: To observe two types ofsecondary complexes, we further devised a binding system using unlabeledBPDE-90-mer to compete with the labeled TMR-BPDE-16-mer for a limitingamount of the antibody. In the absence of the 90-mer, the antibody formstwo complexes with the 16mer*; Ab(16mer*) and Ab(16mer*)₂. In thepresence of BPDE-90-mer, the BPDE-90-mer competes with the 16mer* forthe antibody binding sites. Several complexes containing the 90-mer maybe formed, including Ab(90mer), Ab(90mer)₂, and Ab(16mer*)(90mer). Amongthese complexes, only Ab(16mer*)(90mer) is fluorescent and can bedetected. The observation of Ab(16mer*)(90mer) and the accompanyingdecrease of the Ab(16mer*)₂ in the presence of the 90-mer clearlydemonstrate the redistribution of the two ligands and the bindingstoichiometry. The redistribution of ligands between two binding sitesof the antibody cannot be observed in traditional binding assays thatare based on measurement of total bound ligands.

[0282] In this study, we demonstrated that the DNA adduct and theantibody formed two complexes with the 1:1 and 2:1 stoichiometry.Binding of the antibody with a mixture of the TMR-labeled and unlabeledBPDE-16-mer showed a typical competitive binding behavior when a highconcentration of the unlabeled BPDE-16-mer was used. With a lowerconcentration of the unlabeled BPDE-16-mer, the TMR labeled BPDE-16-merwas partially distributed into the secondary complex. The resultssuggest that the two binding sites of the mouse monoclonal antibody aredependent upon each other, and that the primary binding has a higheraffinity than the secondary binding.

[0283] Scatchard plot is commonly used to characterize molecular bindingevents (Scatchard, 1949). For polyclonal antibody, Scatchard plot iscurved because of the heterogeneous population of the antibodymolecules. For monoclonal antibody, Scatchard plot is considered linearbecause monoclonal antibody is homogeneous with the same affinity andhas two identical antigen-binding sites. However, this does not takeinto account the differences between the secondary and the primarybinding. When the secondary binding is affected by primary binding, theScatchard plot could deviate from linearity. In fact several authorshave observed non-linear Scatchard plots for monoclonal antibodyalthough reasons for the non-linearity was not discussed. It is possiblethat secondary binding and the redistribution may contribute to thenon-linear nature of calibration curves for traditional competitiveassays where the primary and secondary complexes are not separated. Intraditional competitive inmmunoassay, the dose-response curve usually issigmoid in shape with two asymptotes and one point of inflection (Nix,et al., 2000).

We claim:
 1. A method for detecting a binding factor for a probe,comprising: (a) labeling the probe with a fluorophore; (b) incubatingthe labeled probe with a factor or a group of factors which may bind thelabeled probe to form a binding complex; (c) separating the bindingcomplex and the free probe into different fractions; and (d) subjectingeach fraction from step (c) to fluorescence polarization measurementunder conditions wherein the binding complex produces a fluorescencepattern different from that of the free probe, thereby allowingdetection of the binding complex.
 2. The method of claim 1 wherein thefree probe and the complex are separated by using capillaryelectrophoresis.
 3. The method of claim 1 wherein the group of factorscomprises a chemical compound library.
 4. The method of claim 4 whereinthe chemical compound library is a combinatorial library.
 5. The methodof claim 1 wherein the group of factors comprises a mixture of naturalproducts.
 6. The method of claim 6 wherein the mixture of naturalproducts comprises a cell lysate.
 7. The method of claim 1 wherein thegroup of factors comprises nucleic acid.
 8. The method of claim 7wherein the nucleic acid is genomic DNA.
 9. The method of claim 8wherein the probe is capable of binding to modified DNA.
 10. The methodof claim 10 wherein the modified DNA is a DNA adduct.
 11. The method ofclaim 1 wherein the probe is selected from the group consisting ofprotein and nucleic acid.
 12. The method of claim 1 wherein the probehas a molecular weight of less than about 10,000 daltons.
 13. The methodof claim 1 wherein the probe has a molecular weight of less than about5,000 daltons.
 14. The method of claim 1 wherein the probe has amolecular weight of less than about 3,000 daltons.
 15. The method ofclaim 1 further comprising the step of determining binding affinityand/or stoichiometry between the probe and the binding factor.
 16. Themethod of claim 1 wherein the fluorophore is fluorescein.
 17. A methodfor detecting a nucleic acid damage in a nucleic acid sample,comprising: (a) incubating the sample with (i) a polypeptide which iscapable of binding the damaged nucleic acid; and (ii) afluorophore-labeled probe which is capable of forming a complex with thepolypeptide to compete with the damaged nucleic acid for thepolypeptide; and (b) analyzing the incubation mixture under conditionswherein the complex formed between the probe and the polypeptideproduces a fluorescence pattern different from that of a free probe. 18.The method of claim 17 wherein the polypeptide is an antibody which iscapable of binding damaged DNA.
 19. The method of claim 17 wherein theDNA damage is a covalent modification.
 20. The method of claim 17wherein the DNA damage is a benzopyrene addition.
 21. The method ofclaim 17 wherein the DNA sample is genomic DNA.
 22. A method fordetecting a fluorophore labeled probe, comprising: (a) incubating aprobe with a fluorophore under conditions which allow labeling of theprobe by the fluorophore; and (b) subjecting the incubation mixture tofluorescence polarization under conditions wherein the fluorophorelabeled probe produces a fluorescence pattern which is different fromthat of a free probe which is not labeled by the fluorophore.
 23. Themethod of claim 22 further comprising the step of fractionating theincubation mixture.